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STUDIES ON SPERMATOGENESIS AND APOPTOSIS IN THE BOVINE 



By 
VICTOR HUGO MONTERROSO PEREZ 



A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL 

OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT 

OF THE REQUIREMENTS FOR THE DEGREE OF 

DOCTOR OF PHILOSOPHY 

UNIVERSITY OF FLORIDA 

1998 



Copyright 1998 

by 

Victor Hugo Monterroso Perez 



Esta disertacion esta dedicada al Senor Dios todo poderoso, quien creo los cielos y 
la tierra, Dios de Israel, Jehova de los ejercitos. Dios que me infunde aliento, quien 
nunca me ha dejado, quien durante el dia me ha guiado con su nube y con su luz durante 
la noche, quien calma la mar durante la tormenta y el que me lleva a puerto seguro. Al 
Dios que me ha permitido caminar en seco en medio de la mar y en camino donde no lo 
ha habido, El que ha entregado al enemigo en mis manos, El que ha aderezado meza 
delante de mi, El que ha entregado esta victoria en mis manos, El que me ha permitido 
alcanzar esta meta, a quien pertenece toda gloria y honra, a quien pertenece el 
conocimiento y la sabiduria, al Dios de mi salvation. 

- a mi esposa: 

Norma, mujer a quien amo y la companera idonea con la que Dios me ha 
bendecido. 

- a mi hijo: 

Victor Alejandro, quien ha enternecido y ha bendecido mi vida, el que al 
amarle me ha permitido tener una idea del gran amor de Dios para con sus 
hijos. 

- a mis padres: 

Nehemias Monterroso y Lidia Perez de Monterroso, porque de nino me 
instruyeron en los caminos de Dios y por el amor incondicional que me 
han dado a mi, a Norma y a Victor Alejandro. 

- a mis abuelos: 

Victor Manuel Monterroso 1 , Carmen Viuda de Monterroso (Abuelita 
Carmen), Teodoro Alejandro Perez f (Papito Lolo) y Socorro Mazariegos 



de Perez f (Mamita Coco); porque instruyeron a mis padres en los caminos 

de Dios, lo cual me permitio crecer en un hogar temeroso de Dios. 
a mis hermanos: 

Edgar Nehemias y Marisol, Monica Beatriz y Oscar y Lidia Carolina, 
a mis sobrinos: 

Guillermo Javier, Edgar Andres, Jose Pablo, Juan Daniel, Oscar Emilio y 

Benjamin, 
a mis tias: 

Tia Mery y Tia Cori. 
a mis tios: 

Antonio (tio Tono), Ezequiel (tio Cheque) y Lilli*, Alfredo (tio Alfre), 

Jaime (tio Canchito), Waldemar ( tio Walde), y David (tio Davi). 
a mis primos: 

Mireyita, Carlitos, Karinita, Paquito, Michelle, Raul (mi primo 

consentido), Carlos (Humber), Tono, Jorge, Isabel y a todos los que no 

tengo espacio para mencionar. 
a mi familia en los Estados Unidos: 

Norma y Miguel Diaz; Maggie y Marcos Sequeira; Roberto, Priscila, 

Leslie y Lylli Silva; Salvador, Bruni y Salvador Alejandro Robles. 
a la memoria de: 

Gil Mora f , padre de mi esposa. 



al Institute) Evangelico "America Latina": 

Porque en el colegio, siendo nino, se me inculco, que "El principio de la 
sabiduria es el temor a Jehova" (Provervios 1 :7), ahora de adulto, puedo 
reconocer, que "Jehova da la sabiduria, Y de su boca viene el 
conocimiento y la inteligencia" (Provervios 2:6), y que cuando la sabiduria 
entrare en mi corazon, y la ciencia fuere grata a mi alma, la discretion me 
guardara; me preservara la inteligencia, para librarme del mal camino, de 
los hombres que hablan perversidades, que dejan los caminos derechos, 
para andar por sendas tenebrosas; que se alegran haciendo el mal, que se 
huelgan en las pervesidades del vicio; cuyas veredas son torcidas, y 
torcidos sus caminos (Provervios 2:10-15). Porque fue el centro de 
estudios donde aprendi a leer, a escribir, a contar, del Popol Vuh y de la 
Mansion del Pajaro Serpiente. Porque ha sido parte de mi familia en 
nuestras alegrias, en nuestras tristezas, en nuestros logros y en nuestros 
fracasos, y ademas, nos ha visto crecer por tres generaciones (mis padres, 
nosotros y ahora nuestros hijos). Con especialidad al Dr Virgilio Zapata y 
a su esposa, dona Beatriz de Zapata, por su carino hacia nosotros (los 
Monterroso Perez), y a mi maestra, Delfina Herrera de Vega (Seno 
Delfina). 

■ al Instituto "Adolfo V. Hall": 

Porque me instruyeron bajo los codigos del honor y la ciencia, y bajo la 
disciplina militar. Donde aprendi a estudiar para dignificarme, para mas 
tarde honrar a la patria, y a no darme por vencido ni aiin vencido, donde 






quedaron innumrables gotas de mi sudor y de mis lagrimas, donde quedo 
mi adolescencia, y donde aprendi que la patria se llama Guatemala! 
Guatemal! Guatemala!. 

a mi pueblo: 

San Pedro Yepocapa, tierra de mi ninez, tierra donde pude oir la voz de 
Dios en los potentes truenos, tierra donde pude ver la creation de sus 
manos en sus montanas, tierra donde pude sentir el poder de Dios en el 
estruendo y en el poder de su volcan (volcan de Fuego), tierra donde pude 
contemplar la grandeza de Dios al ver sus miles de estrellas, tierra donde 
pude sentir la presencia de Dios en sus vientos fuertes, tierra como en 
Horeb, donde brota el agua de la pena, tierra de cafetos y tierra donde esta 
papito Lolo. 

■ a mi patria: 

Guatemala, pais de la eterna primavera, tierra del quetzal, de la marimba y 
de hombres de maiz. Nation que se ha estremecido, sangrado y llorado 
por la guerra entre hermanos. Y que como resultado, su pueblo ha 
buscado la prescencia de Dios. 



Victor Hugo Monterroso Perez 

Deciembredel998 

Gainesville, Florida, USA 



ACKNOWLEDGMENTS 

I would like to give God all the honor and the glory for this achievement. Also, I 
would like to thank God for allowing me to reach this goal in my life. I thank my wife 
Norma and my son Victor Alejandro for their love and support during all these years of 
graduate studies, making this degree their achievement too. I thank my parents, 
Nehemias y Lidia Monterroso; my brothers and sisters, Edgar and Marisol, Carolina, 
Monica and Oscar; my aunts, Mery and Cori; my family-in-law, Norma and Miguel, 
Maggie and Marcos; my cousins and friends, Roberto and Priscila; my family in 
Gainesville, Salvador and Bruni; my local church, First Assembly of God; and my 
Guatemalan church, Fraternidad Cristiana de Guatemala. 

I want to thank the members of my supervisory committee, Dr. Louis F. Archibald, 
Dr. Claus D. Buerguelt, Dr. Chadwick C. Chase , Jr., Dr. Peter J. Chenoweth, and Dr. 
Kenneth C. Drury. I thank everybody at the USDA/STARS experimental station at 
Brooksville, FL, for all their help in my research. I thank Marty Johnson, Patricia Lewis, 
and Dr. Chi-Chung Chou for their valuable help in my lab work. My gratitude is 
expressed to everybody in RAMS (Dr. Owen Rae, Dr. Carlos Risco, Dr. Art Donovan, 
Dr. Thang Tran, Dr. Jorge Hernandez, Delores Foreman, Fred Bennet, and Jesse Elliot) 
for their support and friendship. Also, I thank my friends: Dr. Carlos Arechiga, Dr. 
Rafael Roman, Dr. Pedro Melendez, Dr. Jose Hugo Urdaz, Dr. Martin Giangreco, Dr. 

vii 



Julian Bartolome, Dr. Antonio Landaeta, Dr. John Crews, Dr. Sandra Bortnik, Dr. Gina 
DeChant, Dr. Lara Maxwell, and Dr. Frank Bernard. 

I want to thank Dr. Charles Courtney, Associate Dean and Sally O'Conell, 
Secretary of the Office for Research and Graduate Studies, Dr. John Harvey, Chair of the 
Department of Physiological Sciences, and Dr. Eleanor Green, Chair of the Department 
of Large Animal Clinical Sciences of the College of Veterinary Medicine at the 
University of Florida for their help and financial support. Also, I would like to thank Dr. 
Alistair Webb, Dr. Richard Johnson, and Dr. Kevin Anderson for the opportunity to work 
with them as a teaching assistant in their courses. 






vni 






TABLE OF CONTENTS 



ACKNOWLEDGMENTS vii 

LIST OF TABLES xii 

LIST OF FIGURES xiv 

ABBREVIATIONS xvi 

ABSTRACT xviii 

CHAPTERS 

1 INTRODUCTION 1 

2 REVIEW OF LITERATURE 6 

Causes of Bull Infertility 6 

Causes of Spermatogenic Dysfunction 7 

Effect of Stressors on Spermatogenesis 7 

Effect of Elevated Temperatures on Spermatogenesis 8 

Effect of Disease on Spermatogenesis 10 

Effect of Toxicity on Spermatogenesis 11 

Effect of Free Radicals on Spermatogenesis 13 

Effect of Hormonal Changes on Spermatogenesis 14 

Spontaneous Degeneration or Germ Cell Death 16 

Effect of Gossypol on Reproduction 17 

Characteristics of Apoptosis or Programmed Cell Death 18 

Characteristics of Necrosis 19 

Genes Reported to Play a Role in the Regulation of Apoptosis 19 

Detection of Apoptosis in Testicular Tissue 21 

Apoptosis in Testicular Tissue 21 

Germ Cell Types Involved in Apoptosis 23 



IX 



Heat Shock Proteins 24 

Heat Shock Proteins in Testicular Tissue 24 

Scrotal Insulation Model 26 

Effect of E. coli Endotoxin Infusion 28 

Effect of Elevated Temperatures on Testosterone Levels 29 

Effect of Elevated Temperatures on Gonadotropins 31 

Breed Differences in Susceptibility to Elevated Temperatures 32 

3 ATTEMPTS TO INDUCE AND ASSESS SPERMATOGENIC APOPTOSIS IN 
BULLS 37 

Introduction 37 

Materials and Methods 39 

Materials 39 

Experiment 1 39 

Experiment 2 44 

Statistical Analysis 45 

Results 46 

Experiment 1 46 

Experiment 2 59 

Discussion 73 

4 EFFECT OF DIETS CONTAINING FREE GOSSYPOL AND VITAMIN E ON 
SPERMATOGENESIS AND SPERMATOGENIC APOPTOSIS IN YOUNG 
HOLSTEIN BULLS 80 

Introduction 80 

Materials and Methods 81 

Materials 81 

Experimental design 82 

Statistical Analysis 88 

Results 88 

Discussion 91 

5 BREED EFFECTS ON SPERMATOGENESIS AND SPERMATOGENIC 
APOPTOSIS IN BULLS 96 

Introduction 96 

Materials and Methods 98 

Materials 98 

Experiment 1 99 

Experiment 2 100 

Experiment 3 1 02 






Statistical Analysis 102 

Results 103 

Experiment 1 103 

Experiment 2 103 

Experiment 3 107 

Discussion Ill 

6 SUMMARY AND CONCLUSIONS 118 

LIST OF REFERENCES 127 

BIOGRAPHICAL SKETCH 148 



XI 



LIST OF TABLES 

Table page 

2-1. Concentration of Hsp 70 in bovine tissue as determine by ELISA 25 

3-1 . Effect of treatment (control, endotoxin, or scrotal insulation) on sperm motility, 

normal spermatozoa, primary abnormalities, and secondary abnormalities .... 51 

3-2. Semen and testicular traits at 8 d in control, E. coU endotoxin or scrotal insulated 
(SI) treated bulls 52 

3-3. Sertoli cells, germ cells (spermatogonia A, preleptotene, pachytene, and round 

spermatid 8), and Sertoli:germ cells ratio in 20 seminiferous tubules stage VIII in 
control, E. coH endotoxin infusion (100 ng/kg) (endotoxin), or scrotal insulated 
for 48 h (SI) bulls 53 

3-4. Sperm motility, normal spermatozoa, primary abnormalities, and secondary 

abnormalities in Angus, Romosinuano, and Senepol 60 

3-5. Body weight, rectal, and scrotal temperatures in controls and treated bulls of 3 

breeds at 24 h following initiation of scrotal insulation (SI) in bulls 64 

3-6. Semen and testicular traits in control or 48 h scrotal insulated (SI) bulls 66 

3-7. Semen and testicular traits in Angus, Romosinuano, and Senepol bulls at 24 h 

following initiation of scrotal insulation (SI) 67 

3-8. Heat shock protein 70 (jxg/ml) in semen of Angus, Romosinuano, and Senepol 

bulls 68 

4-1. Initial and final composition of dietary supplements 83 

4-2. Average composition of dietary supplements 84 

4-3. Sperm morphology and testicular traits in young Holstein bulls fed diets 

containing gossypol, gossypol plus vitamin E, and controls 89 



xn 



5-1. Physical and testicular traits in young B. taurus and B indicus bulls 104 

5-2. Spermatogenic traits and extragonadal sperm reserves in young B. taurus and B 

indicus bulls 105 

5-3. Physical and testicular traits in young crossbred bulls 106 

5-4. Spermatogenic traits and spermatogenic apoptosis in young crossbred bulls . . 108 

5-5. Testicular traits, spermatogenesis, and spermatogenic apoptosis in Brahman and 
miniature Brahman bulls 110 

5-6. Testicular traits, spermatogenesis, and spermatogenic apoptosis in Romosinuano 
bulls 112 



xin 



LIST OF FIGURES 



Figure Page 

3 - 1 . Effect of E.coli endotoxin infusion ( 1 00 ng/kg) on respiration rate, heart rate, and 
rectal temperature in bulls 48 

3-2. Spermatozoal motility in control, E. coli endotoxin infused (100 ng/kg), or 

scrotal insulated for 48 h (SI) 49 

3-3. Percentage of normal spermatozoa, primary abnormalities, and secondary 

abnormalities in control, E.coli endotoxin infused (100 ng/kg) (endotoxin), or 
scrotal insulated for 48 h (SI) bulls 50 

3-4. Sertoli cells, germ cells (spermatogonia A, preleptotene, pachytene, and round 

spermatid 8), and Sertoli:germ cells ratio in 20 seminiferous tubules stage VIII in 
control, R coli endotoxin infusion (100 ng/kg) (endotoxin), or scrotal insulated 
for 48 h (SI) bulls 54 

3-5. Apoptotic cells per stage VIII seminiferous tubule in control, E. coh endotoxin 

infusion (100 ng/kg) (endotoxin), or scrotal insulated for 48 h (SI) bulls 55 

3-6. Apoptotic cells per seminiferous tubule in control, R. coh endotoxin, and 48 h of 
scrotal insulation (SI). Spermatogenic apoptosis assessed by Tunel staining ... 56 

3-7. Effect of testicular concentration on light absorbance at 405 nm, using a cell 

detection ELISA 57 

3-8. Effect of E.coli endotoxin infusion (100 ng/kg) or 48 h of scrotal insulation(SI) 
on spermatogenic apoptosis, DSPG (daily sperm production/g) and DSP (daily 
sperm production) 5g 

3-9. Spermatozoal motility in control or scrotal insulated for 48 h (SI) Angus, 

Romosinuano, and Senepol bulls 61 



xiv 



3-10. Percentage of normal spermatozoa, primary abnormalities, and secondary 

abnormalities in control or scrotal insulated for 48 h (SI) 62 

3-11. Percentage of normal spermatozoa, primary abnormalities, and secondary 

abnormalities in Angus, Romosinuano, and Senepol bulls 63 

3-12. Heat shock protein 70 (Hsp70, /ug/ml) on semen over time in control or scrotal 

insulation ( SI 24 h) 69 

3-13. Heat shock protein 70 (Hsp70, Mg/ml) on semen over time, regardless of 

treatment, in Angus, Romosinuano, and Senepol bulls 70 

3-14. Heat shock protein 70 (Hsp70, //g/ml) on semen over time, regardless of breed 

and treatment 71 

3-15. Effect of 48 h of scrotal insulation (SI) on apoptotic cells per seminiferous tubule 
in tissue harvested immediately following (d 2) removal of scrotal insulation, or 2 
days later (d 4). Spermatogenic apoptosis assessed by Tunel stain 72 

3-16. Effect of 48 h of scrotal insulation (SI) on spermatogenic apoptosis (absorbance 
at 405 nm), daily sperm production/g (DSPG), and daily sperm production (DSP) 
in tissue harvested immediately following (d 2) removal of scrotal insulation, or 2 
days later (d 4) 74 

3-17. Spermatogenic apoptosis (absorbance at 405 nm), daily sperm production/g 

(DSPG), and daily sperm production (DSP) in Angus, Romosinuano, and Senepol 
bulls 75 

4-1 . Apoptotic cells per seminiferous tubule in bulls fed gossypol (GOSS), gossypol 
plus vitamin E (G+VITE), and controls (CONT). Spermatogenic apoptosis, 
assessed by Tunel staining 90 

4-2. Levels of spermatogenic apoptosis and sperm production in bulls fed gossypol 

(GOSS), gossypol plus vitamin E (G+VITE), and controls (CONT) 92 

5-1 . Spermatogenic apoptosis in crossbred bulls (SA= Senepol x Angus, TA= Tuli x 

Angus, and BA= Brahman x Angus) 109 



xv 



ABBREVIATIONS 



AI 


Artificial insemination 


ANOVA 


Analysis of variance 


BA 


Brahman x Angus 


BSE 


Breeding soundness evaluation 


B. taurus 


Bos taurus 


B. indicus 


Bos indicus 


BW 


Body weight 


°C 


Celsius degrees 


Ca 


Calcium 


CP 


Crude protein 


CSM 


Cottonseed meal 


Cu 


Copper 


DM 


Dry matter 


DNA 


Deoxyribonucleic acid 


DPBS 


Dulbecco's Phosphate Buffered Saline 


DSP 


Daily sperm production 


DSPG 


Daily sperm production per g 


EDTA 


Ethylenediaminetetraacetic acid 


FSH 


Follicle-stimulating hormone 


GnRH 


Gonadotropin releasing hormone 


GSH 


Reduced glutathione 


hCG 


human chorionic gonadotropin 


Hsp 


Heat shock protein 


IL-6 


Interleukin-6 


IV 


Intravenous 


IVF 


In vitro fertilization 


K 


Potassium 


LH 


Luteinizing hormone 


LSMeans 


Least squares means 


MDA 


Malondiadldehyde 


Mg 


Magnesium 


MM 


microtiterplate-modules 


Mw 


Molecular weight 


N 


North 


•OH 


Hydroxyl radical 



XVI 



p 


Phosphorus 


PGFlcc 


Prostaglandin F-l alpha 


ppm 


Parts per million 


ROS 


Reactive oxygen species 


SA 


Senepol x Angus 


SBM 


Soybean meal 


SDS-PAGE 


Sodium dodecyl sulfate polyacrylamide gel electrophoresis 


SEM 


Standard error of the mean 


SI 


Scrotal insulation 


TA 


Tuli x Angus 


TD 


Time divisor 


TdT 


terminal deoxynucleotidil transferase 


TMR 


Total mixed ration 


TNF 


Tumor necrosis factor 


vs 


Versus 


v:v 


Volume:volume 


W 


West 


w:v 


Weight:volume 



UNITS 



u 


Units 


ID 


International unit 


kg 


Kilogram 


g 


Gram 


mg 


Milligram 


Hg 


Microgram 


1 


Liter 


M 


Molar 


ml 


Milliliter 


Ml 


Microliter 


mU 


Milliunits 


mo 


Month 


wk 


Week 


d 


Day 


h 


Hour 


min 


Minutes 



xvn 



Abstract of Dissertation Presented to the Graduate School 

of the University of Florida in Partial Fulfillment of the 

Requirements for the Degree of Doctor of Philosophy 



STUDIES ON SPERMATOGENESIS AND APOPTOSIS IN THE BOVINE 

By 

Victor Hugo Monterroso Perez 

December, 1998 

Chairperson: Peter J. Chenoweth 

Major Department: Veterinary Medical Sciences 

The main objective was to study the effect of elevated temperatures, gossypol, 
Vitamin E, and breed differences on spermatogenesis and spermatogenic apoptosis. First, 
induction and assessment of apoptosis were studied. Experiment (exp.) 1 (Angus, n=l 1), 
included; 1) control, 2) E. coli endotoxin, and 3) scrotal insulation (SI). Spermatogenesis 
(daily sperm production, DSP and DSP/g, DSPG) and apoptosis were assessed at 10 d. 
Experiment 2, (Angus, An; Senepol, Se; Romosinuano, Ro; n=18) included; 1) control 
and 2) SI. Assessment was done at 2 d and 4 d. In exp. 1 and 2 treatment had no effect 
on spermatogenesis and apoptosis. 

Second, effects of gossypol and vitamin E on spermatogenesis and apoptosis were 
tested. Young Holstein bulls (n=24) were supplemented with either; 1) CONT (soybean 
meal, corn, and vitamin E, 2) GOSS (cotton seed meal, CSM, corn, and vitamin E), and 

xviii 



3) G+VITE (CSM, corn, and vitamin E). Spermatogenesis was less (P<0.05) in GOSS 
than CONT or G+VITE. Gossypol had no effect on apoptosis, although there was an 
inverse relationship between spermatogenesis and apoptosis (PO.05). 

Third, young B. taurus, B_. indicus, and crossbred bulls were evaluated during the 
summer in Florida. In exp. 1 (n=36), An, Ro, Brahman (Br), and Nelore x Brahman bulls 
were used. Breed had no effect on spermatogenesis, extragonadal sperm reserves (ESR) 
(caput), and ESR (cauda). However, ESR (corpus) in Br was higher than in An and Ro 
(P^ 0.05). In exp. 2, crossbred bulls (n=l 12; Senepol x Angus, SA; Tuli x Angus, TA; 
Brahman x Angus, BA) were used, and DSP (perhaps DSPG) was less in BA than in SA 
and TA (PO.05). In exp. 3, mature Brahman (n=2) and miniature Brahman (n=5) bulls 
were used, and DSP was less in miniature than in Brahman bulls (PO.05). In exp. 2 and 
3 breed did not affect apoptosis. 

This study failed to observed increases in spermatogenic apoptosis as a result of 
E. coli endotoxin, SI, gossypol, summer temperatures, and breed. In general, low levels 
of apoptosis were detected and reasons for this may include inappropriate sampling times 
and test sensitivity. 



xix 



CHAPTER 1 
INTRODUCTION 



Overall calving rates in beef cattle in the Gulf states are estimated at between 70 
and 75% (Florida Agriculture Statistics, 1985-1993); a figure well below optimum. 
Increasing reproductive efficiency in a cost-effective way, would increase efficiency of 
beef production, profitability, and decrease pressures upon scarce natural resources. Most 
beef cattle in the US use natural breeding where the bull plays a pivotal reproductive role. 
In turn, the reproductive efficiency of bulls is related to their spermatogenic capabilities 
(Chenoweth, 1994a). Elevated environmental temperatures can impair spermatogenesis 
in bulls which leads to decreased semen quality with associated lowered fertility (Casady 
et al., 1953; Skinner and Louw, 1966; Rhynes and Ewing, 1973; Meyerhoeffer et al., 
1985; Ax et al., 1987). In addition, bull spermatogenesis has been reported to be 
adversely affected by elevated body temperatures, toxins, and viral disease (Casady et al., 
1953; Austin et al., 1961; Johnston et al., 1963; Bartak, 1973; Burgess and Chenoweth, 
1975; Larsen and Chenoweth, 1990; Vogler et al., 1993; Chenoweth et al., 1994). 
Common semen characteristics in affected bulls include lower sperm numbers, lower 
motility, increased abnormal sperm, and increased dead sperm, all of which have been 
associated with infertility (Saacke, 1982). Breed differences occur in susceptibility to 
heat stress, with less adverse reproductive effects being reported in males of B. indicus 



breeds compared to those of B. taurus derivation (Chenoweth, 1991). In addition to the 
adverse reproductive effects attributable to specific stressful events, sperm output is 
generally lower than expected from theoretical extrapolation of spermatogenic precursors 
in the mouse, rat, and human with such findings being associated with germ cell 
degeneration and depletion (Oakberg, 1956; Russel and Clermont, 1977; Johnson et al., 
1984). In the rat, the cells most implicated in this degenerative process include 
spermatogonia (Kerr, 1992) and spermatocytes (Brinkworth et al., 1995). 

Lowered sperm output is usually associated with degeneration of the 
spermatogenic epithelium, a process in which apoptosis (or programmed cell death) 
probably plays an important role. Apoptosis has been observed and studied in a variety of 
cells and tissues including those of the developing embryo, regressing corpus luteum, 
lymph nodes, gastrointestinal tract, tumors, and testes (Wyllie et al., 1980; Bowen and 
Bowen, 1990; Miething, 1992; Palumbo and Yeh, 1994). In structural terms, apoptosis is 
characterized by condensation of chromatin and cytoplasm, breakage of DNA in a 
characteristic ladder pattern on electrophoresis, and the formation of apoptotic bodies 
which are subsequently phagocytosed by macrophages or Sertoli cells in testes. 
Apoptosis has been observed in the testes of guinea pigs, rats, hamsters, deer, cattle, and 
other mammals (Allan et al., 1992; Shikone et al., 1994; Higst and Blottner, 1995). 
Testicular apoptosis in rats has been induced by elevated temperatures or by withdrawal 
of GnRH, LH, FSH, and testosterone (Tapanainen et al., 1993; Shikone et al., 1994). It 
has also been associated with an increase of diadem/crater defects in ejaculated sperm, 
and observed during seasonal testicular regression in the roe deer (Hingst and Blottner, 



1995). Increased diadem/crater defects of ejaculated sperm have also been associated 
with elevated body temperature (Malmgren and Larsson, 1985), scrotal insulation (Vogler 
et al., 1993) and spermatoxic agents such as ethylene dibromide (Courtens et al., 1980; 
Eljack and Hrudka, 1979). Such defects are first detectable by microscopy in round 
spermatids (Larsen and Chenoweth, 1990). 

A common mechanism underlying these different disruptions to spermatogenesis 
may only be hypothesized at present. However, it is well established that an imbalance 
between the generation of reactive oxygen species (ROS) and their elimination can result 
in cellular damage (Cagnon et al., 1992). Oxidative stress has been reported to induce 
apoptosis in thymocytes and embryonic cortical neurons (Ratan et al., 1994; Wolfe et al., 
1994). Excessive free radical formation (or lack of elimination) has also been proposed 
as a major cause of cell damage resulting from heat shock (Loven, 1988). Free radicals 
can react with lipids to form toxic by-products such as lipid hydroperoxidase, epoxides, 
and aldehydes that react with other lipids to cause membrane lipid peroxidation and 
disruption of membrane integrity (Savanian and Hochstein, 1985; Halliwell, 1987; Radi 
et al., 1991). Free radicals can also react with proteins and purine and pyrimidine bases 
to cause proteolysis and oxidation (Salo et al., 1990; Fraga et al., 1990). Sperm contains 
enzymes implicated in the neutralization of ROS, and damage to sperm occurs when the 
equilibrium is disturbed between the amounts of ROS produced and the scavenging 
mechanisms, leading to lower sperm motility and viability as well as damage to the 
axoneme (Cagnon et al., 1992). 



Damage to the spermatogenic epithelium of bulls caused by gossypol, a free 
radical inducer and sperm axoneme disrupting agent, has been effectively countered with 
antioxidant administration in the form of vitamin E (Velasquez-Pereira et al., 1995). 

Recognition of the role of apoptosis in spermatogenic dysfunction introduces new 
possibilities for the diagnosis and possible prevention of male infertility. However, in 
order to use this criterion in the evaluation of spermatogenesis, important prerequisites to 
consider include the establishment of quantifiable links between testicular stress, 
spermatogenic dysfunction, and apoptosis, and the identification of predictable markers in 
ejaculated semen and/or sperm. A potential candidate in this latter group include elevated 
levels of hsp70 in semen. 

The prospect of improving reproductive efficiency in beef herds by boosting bull 
fertility is attractive, particularly if it can be achieved with minimal cost and intervention. 
Bulls often suffer decreased semen quality and sperm output during the hottest months of 
the year with deleterious effects upon both natural and artificial breeding programs. A 
better understanding of the mechanisms involved in spermatogenic dysfunction could 
allow protective mechanisms and strategies to be devised. 

The objective of this work was to determine the effectiveness of testicular 
stressors (testicular insulation, E. coli endotoxin, and gossypol) to induce spermatogenic 
apoptosis and to compare this with summer environmental effects on spermatogenesis in 
Florida. Possible differences between B. taurus and B. indicus genotypes in their 
susceptibility to environmentally induced spermatogenic apoptosis and efficiency were 
investigated. To achieve these objectives, methods of identifying and quantifying 



spermatogenic apoptosis were compared with quantitative estimates of spermatogenesis. 
Methods of alleviating or preventing stress effects on bull spermatogenesis, such as the 
use of antioxidants were investigated. Two general hypotheses were tested in this study; 
first, that stressors will increase spermatogenic apoptosis in treated bulls, and second, that 
spermatogenic apoptosis will be lower in tropically adapted than in temperate breeds 
during periods of heat stress. 



CHAPTER 2 
REVIEW OF LITERATURE 



Causes of Bull Infertility 



Infertility in the bull may be caused by a variety of factors including genetic 
problems, cryptorchidism, elevated temperatures, disease, toxicity, and others (Andersson 
et al., 1990; Larsen and Chenoweth, 1990; Chenoweth et al., 1994; Marcus et al., 1997; 
Steffen, 1997). 

Genetic or congenital problems reported to cause infertility in the bull have 
included chromosomal abnormalities, spermatozoal abnormalities, cryptorchidism, 
inguinal hernia, testicular hypoplasia, segmental aplasia/hypoplasia of the Wolffian duct 
system, persistent penile frenulum, as well as others (Andersson et al., 1990; Mcfeely et 
al., 1993; Chenoweth et al., 1994; Marcus et al., 1997; Steffen, 1997). Also, elevated 
temperatures, both environmental and experimental have been reported to induce 
infertility in the bull (Casady et al., 1953; Johnston and Branton, 1953; Johnston et al., 
1963; Sidibe et al., 1992; Vogler et al., 1993). Diseases such as ephemeral fever, 
eperythrozoonosis caused by Eperythrozoon wenyonii infection, and Haemophilus 
somnus infection have been associated with bull infertility (Burgess and Chenoweth, 
1975; Barber et al., 1994; Montes et al., 1994). Toxicity has also been associated with 
infertility by causing spermatozoal abnormalities, decreasing sperm production, or a 



7 
combination of both. For example, administration of dexamethasone or ethylene 
dibromide has been associated with crater defects of sperm (Coulter, 1976; Eljack and 
Hrudka, 1979; Courtens et al., 1980). Another toxic agent associated with bull infertility 
is gossypol, a component of cotton seed which has been associated with increased 
spermatozoal malformations and diminution of spermatogenesis (Chase et al., 1994; 
Chenoweth et al., 1994; Velasquez-Pereira et al., 1995). 

Causes of Spermatogenic Dysfunction 

Effect of Stressors on Spermatogenesis 

In natural breeding systems the bull plays a pivotal role in beef production. 
Suboptimal bull fertility undoubtedly contributes significantly to the relatively low calf 
crop reported for Gulf States such as Florida (Florida Agricultural statistics, 1985-1993). 
A number of stressors can adversely affect bull fertility, including elevated temperatures 
and disease. High ambient temperatures can compromise spermatogenesis in the bull. 
These adverse effects are often associated with semen changes including lowered sperm 
concentration, motility and membrane integrity, as well as increased morphological 
abnormalities. All of these effects have been associated with lowered fertility (Casady et 
al., 1953; Johnston et al., 1963; Ax et al, 1987; Wolfe et al., 1993). The spermatogenic 
ephitelium responds to stressors in a stereotyped manner, with disruption of chromatin 
condensation being a common manifestation. This in turn can lead to the diadem/crater 
defect in ejaculated sperm, for which earliest evidence may be found in round spermatids 
(Larsen and Chenoweth, 1990). This defect is considered to represent a morphological 



8 
marker for a wide variety of testicular insults, having been induced by scrotal insulation 
(Vogler et al., 1993), ethylene dibromide toxicity (Courtens et al., 1980), seasonal 
changes (Haigh et al., 1984), and administration of dexamethasone (Coulter, 1976). 
Another important morphological marker commonly associated with spermatogenic 
damage is disruption of the sperm midpiece, varying from subtle gaps and discontinuities 
to more bizarre manifestations. Such midpiece abnormalities may be observed following 
gossypol-induced spermatogenic damage in bulls (Chenoweth et al., 1994). 
Effect of Elevated Temperatures on Spermatogenesis 

Disruption or diminution of spermatogenesis in bulls has been induced either 
experimentally or naturally by unilateral castration, diseases such as ephemeral fever, or 
by elevated temperatures secondary to scrotal insulation, environmental chambers, and 
cryptorchidism (Bartak, 1973; Burgess and Chenoweth, 1975; Ross and Entwistle, 1979; 
Wolfe etal., 1993). 

The combination of elevated environmental temperatures and high humidity can 
also compromise spermatogenesis in bulls (Johnston and Branton, 1953; Casady et al., 
1953; Johnston et al., 1963). For example, Johnston and Branton (1953) reported that 
fertility of dairy bulls declined during the summer months in Louisiana, and 
spermatogenesis was lowered in Guernsey bulls when bulls were exposed to continuous 
elevated temperatures using environmental chambers (Casady et al., 1953). Also, 
Johnston et al. (1963) exposed purebred Holstein Friesian, Brown Swiss and Red Sindhi 
crossbred bulls to a maximum of 104° F and 54% relative humidity and a minimum of 



82° F and 72% relative humidity for an 8 h period every day for 7 days using 
environmental chambers. Elevated temperature treatment resulted in lower sperm 
motility, concentration, and total number of spermatozoa. Also, an increase in 
spermatozoal abnormalities were observed 4 to 6 weeks after heat stress. When initial 
motility, percentage of abnormal spermatozoa, sperm concentration, and total number of 
spermatozoa, obtained 3 weeks prior to heat stress treatment were compared with values 
obtained 4 to 6 weeks after heat stress treatment, motility declined (P<0.05) from 100 to 
61%, 100 to 81%, 100 to 81%, and 100 to 92% in Holstein, R. Sindhi x Holstein, Br. 
Swiss and R. Sindhi x Br. Swiss, respectively. Spermatozoal concentration declined from 
1 194 to 582, 1239 to 947, 935 to 420, and 1 126 to 789 million spermatozoa/ml for the 
same breeds, respectively. Total number of spermatozoa per ejaculate also declined 
(PO.05) from 6372 to 4927, 13803 to 1 1949, 4998 to 2817, and 10214 to7791 million 
for the same respectively, while the percentage of abnormal spermatozoa increased from 
9 to 33, 1 1 to 24, 16 to 28, and 16 to 18 %, respectively. 

Meyerhoefer et al. (1985) exposed 16 yearling Angus bulls to either control or 
heat stress temperatures using environmental chambers. Heat stress bulls were exposed 
to 35 ° C for 8 h and 3 1 ° C for 1 6 h during an 8 week period, and control bulls were 
maintained at 23 ° C. The heat treatment resulted in lower sperm motility compared with 
controls (P <0.01), and increased percentages of aged acrosomes and abnormal 
spermatozoa both (P< 0.05). 

Austin et al. (1961) divided 12 Hereford bulls into control or scrotal insulation 
(SI) for 48 or 72 h. Scrotal skin temperature was increased by approximately 3 ° F in 






10 
treated bulls compared with control bulls. Also, sperm viability, percent normal, and 
sperm concentration all decreased (65, 60, and 60 %, respectively) in treated bulls 
compared to controls. Scrotal insulation for 48 h caused morphological changes in bull 
spermatozoa (Vogler et al., 1993). In the latter study, abnormalities were first assessed at 
d 12 after SI and peaked at d 18. The predominant abnormalities at different intervals 
from SI were tailless sperm that appeared at d 12-15, diadem defects at d 18, pyriform 
heads and nuclear vacuoles at d 21, knobbed acrosomes at d 27, and "Dag" defects at d 
30. 

Semen quality in bulls has also been reported to decrease during the hottest 
months of the year. In Florida, Fields et al. (1979) reported that semen quality and 
testicular size decreased in Hereford bulls during the summer months. Similar findings 
were reported by Chase et al. (1993) in Angus but not in Senepol bulls. 
Effect of Disease on Spermatogenesis 

Disease processes associated with pyrexia can cause impaired spermatogenesis. 
For example, bovine ephemeral fever, a viral disease of cattle which causes an acute 
pyrexia, has been associated with increased sperm mid-piece abnormalities (Burgess and 
Chenoweth, 1975). In another study, Bartak (1973), reported that unilateral mumps 
orchitis resulted in disruption of spermatogenesis in 50% of human patients, as exhibited 
by increasing oligospermia, necrospermia, and azoospermia. However, it was not readily 
apparent in either of these studies if the disruption of spermatogenesis occurred as a result 
of fever, elevation of cytokines, or due to direct toxic or pathogenic effects. 



11 

Effect of Toxicity on Spermatogenesis 

Compounds such as gossypol, cyclophosphamide, and phthalic acid esters have 
been reported to disrupt spermatogenesis (Randel et al., 1992; Chase et al., 1994; 
Chenoweth et al., 1994; Velasquez-Pereira et al., 1995; Richburg and Boekelheide, 1996; 
Caietal., 1997). 

Gossypol has been shown to have negative effects on male reproduction in 
humans, monkeys, rabbits, hamsters, rats, mice, and bulls (National Coordinating Group 
on Male Antifertility Agents, 1978; Chang et al., 1980; Hahn et al., 1981; Saksena and 
Salmonsen, 1982; Shandilya et al., 1982; Wong et al., 1984; Chase et al., 1994; 
Chenoweth et al., 1994; Velasquez-Pereira et al., 1995). These effects included lowered 
sperm motility and sperm counts, increased sperm malformation, and azoospermia (Liu et 
al., 1987; Risco et al., 1993; Chenoweth et al., 1994). In men, gossypol ingestion at 20 
mg/d for 75 days followed by weekly doses of 50 mg, resulted in sperm concentrations 
lower than 4 million/ml in ejaculates from 99.9% of treated men (National Coordinating 
Group on Male Antifertility agents, 1978). 

When yearling Holstein bulls were fed either 6 or 30 mg total gossypol/kg BW/d 
for 60 or 42 days, no apparent deleterious effects were observed in either semen quality or 
spermatogenesis (Jimenez et al., 1989). In contrast, when postpubertal Brahman bulls 
were fed 8.2 g of free gossypol/day for an 1 1 week period, the percentage of normal 
spermatozoa (49 ± 9.8 vs 83 ± 3.2 %; P=0.001) as well as sperm motility (52 ± 9.8 vs 82 
± 6.2 %; P=0.04) was lower in treated bulls when compared to controls (Risco et al., 



12 
1993). Chase et al. (1994) reported that when Spring-born American Brahman bulls were 
fed 0, 6 or 60 mg/kg BW/d of gossypol from weaning through puberty, bulls fed 60 
mg/kg BW/d reached puberty at an older age than bulls fed or 6 mg/kg BW/d (613 vs 
550 or 528 d; P=0.05). However, no differences in semen quality among groups were 
found. Chenoweth et al. (1994) showed that bulls fed 8.2 g of free gossypol/day during 
an 1 1 week period had lower sperm motility, normal spermatozoa, and sperm production 
(described as daily sperm production and daily sperm production/g of testicular tissue) all 
(PO.05) than control bulls, and the proportion of sperm midpiece abnormalities was 
higher in gossypol treated than in control bulls (P=0.05). In a more recent study, dairy 
bulls fed 14 mg free gossypol/kg BW/d showed a tendency for lower testicular and 
epididymal weights than controls or bulls fed with the same amount of free gossypol plus 
4000 IU Vit E/head/d (245.76 ± 15.58 g vs 262.05 ± 15.58 g and 285.28 g; P=0.1 and 
19.44 ± 1.92 g vs 24.68 ± 1.92 g and 25.97 ± 1.92 g; P<0.1) (Velasquez-Pereira, 1995). 
Other toxic agents such as phthalic acid esters and cyclophosphamide have been 
associated with decreased fertility by inducing spermatogenic dysfunction that resulted in 
increased spermatogenic apoptosis in humans and rats. Phthalic acid esters are found in 
plastizers in food packaging and biomedical devices, and these compounds have been 
associated with low fertility and testicular atrophy in humans (Thomas and Thomas, 
1984; Albro, 1987). Richburg and Boekelheide (1996) treated 28-day-old Fischer rats 
with mono-(2-ethylhexyl) phthalate (MEHP) at a dose of 2 g/kg per os. Rats killed at 0, 
3, 6, or 12 h after treatment showed collapse of Sertoli cell vimentin filaments after 3 
hours of MEHP treatment. Sertoli cell function was adversally affected and this resulted 



13 
in disruption of spermatogenesis as determined by increased spermatogenic apoptosis. 

Cyclophosphamide, a drug used in chemotherapy, has been reported to decrease 
fertility in humans and animals by inducing oligozoospermia or azoospermia (Qureshi et 
al., 1972; Trasler et al., 1986; Watson et al., 1985). Cai et al. (1997), treated male 
Sprague-Dawley rats with cyclophosphamide (70 mg/kg BW). The animals were killed 
and testes collected at 0, 4, 8, 12, 18, 24, and 48 h after treatment. Apoptosis of germ 
cells was observed at all stages of spermatogenesis. However, cell death via apoptosis 
was higher in spermatogonia and spermatocytes in stages I-IV and XI-XIV, suggesting 
that oligozoospermia and azoospermia resulted from cell death of germ cells. 
Effect of Free Radicals on Spermatogenesis 

Oxidative stress has been reported to induce apoptosis in thymocytes and 
embryonic cortical neurons (Ratan et al., 1994; Wolfe et al., 1994). Reactive oxygen 
species (ROS), which contain an unpaired electron, can increase in tissues as a result of 
exposure to elevated temperatures (Freeman et al., 1990). These can react with lipids to 
form toxic by-products such as lipid hydroperoxidase, epoxides, and aldehydes that then 
react with other lipids to cause lipid peroxidation and disruption of the integrity of 
membranes (Savanian and Hochstein, 1985; Halliwell, 1987; Radi et al., 1991). Reactive 
oxygen species can also react with proteins, purine, and pyrimidine bases to cause 
proteolysis and oxidation (Salo et al., 1990; Fraga et al., 1990). Reactive oxygen species 
are proposed as major contributors to cell damage resulting from heat shock (Loven, 
1988). Laskowska-Lita and Szymanska (1989) used malondiadldehyde (MDA) formation 



14 
to measure enzymatic and non-enzymatic lipid peroxidation in bull testis tissue (88.21 ± 
3.01 and 103.2 ± 4.23 nmol/mg protein, respectively). However, when 50 uM of the 
antioxidant glutathione (GSH) was present, non-enzymatic peroxidation MDA 
concentration dropped from 103.3 ± 12 to 11 ± 0.9 nmol/15 minutes per tissue. The 
ability of antioxidants to protect spermatogenic tissue against oxidative damage was 
apparent in a recent study in which the previously established spermatoxic effects of 
gossypol, presumably caused by stimulating free radical production in bulls (Chenoweth 
et al., 1994), were countered by administration of vitamin E at 4000 IU/d (Velasquez- 
Pereira et al., 1995). In this experiment, Velasquez-Pereira (1995), reported that 
antioxidants (4000 IU Vit E/head/d) added to a diet containing 14 mg free gossypol/ kg/ 
body weight, provided a tendency of a protective effect evidenced by increased testicle 
weight (285.28 g in Vitamin E vs 262.05 and 245.67 g, in control and gossypol groups 
respectively; P<0.1). 
Effect of Hormonal Changes on Spermatogenesis 

Pituitary gonadotropins and testosterone are essential for the normal maintenance 
of spermatogenesis (Garner and Hafez, 1993). Sertoli cells, which represent the somatic 
component of the seminiferous epithelium, maintain spermatogenesis under the stimulus 
of FSH and androgens (Parvinen, 1982; Steinberger, 1991; Garner and Hafez, 1993). 
Plasma membrane receptors for FSH and intracellular receptors for androgens are present 
on the Sertoli cell (Means et al., 1976; Heckert and Griswald, 1991; Garner and Hafez, 
1993). Androgens are secreted by Ley dig cells under the influence of LH (Garner and 



15 
Hafez, 1993), and imbalances in either gonadotropins or androgens could disrupt 
spermatogenesis (Tapanainen et al., 1993; Troiano et al., 1994; Blottner et al., 1996; 
Brinkworth et al., 1995; Hikim et al., 1997). 

Rhynes and Ewing (1973), exposed eight Hereford bulls to 21 ° C and 50% 
relative humidity for seven weeks (control period), then four bulls were heat stressed at 
35.5° C and 50% relative humidity for seven weeks. Heat stress increased rectal 
temperature and respiration rate (P<0.01) when compared to controls. Also, 
spermatogenesis as evaluated by semen traits and histology, was impaired by heat 
treatment, and testosterone plasma concentration declined to 43% of control levels 
(P<0.01) during the first two weeks of heat treatment. 

In another study, Sidibe et al. (1992) treated four Swedish Red and White bulls 
with 96 h of scrotal insulation. Heparinized blood was taken every 2 h for a 24 h period 
at two week intervals for testosterone, LH, and Cortisol determination. Testosterone 
levels had a tendency to decrease and LH to increase during the period of severe testicular 
degeneration, whereas the opposite was observed during the period of testicular 
regeneration. Cortisol levels decreased significantly when bulls were analyzed 
individually at 1 and 1 5 weeks after scrotal insulation, suggesting that testicular 
degeneration is associated with changes in testosterone and LH levels. 



16 
Spontaneous Degeneration or Germ Cell Death 

Spontaneous death of testicular germ cells appears to be common in many species 
(Roosen-Runge, 1973; Clermont, 1972). In the absence of overt stress, sperm output is 
often lower than expected in mice, rats, and humans when compared with the theoretical 
number that should be obtained from spermatogonial precursors (Oakberg, 1956; Russell 
and Clermont, 1977; Johnson et al, 1984; Kerr, 1992). The reason for such wastage is not 
known. The contribution of apoptosis to spermatogenic reduction is unknown and may 
only be hypothesized at present, even though apoptosis is often observed in 
spermatogenic tissue. Kerr (1992) described spermatogonial deletion as a cause of 
testicular germ-cell loss in rats, an effect presumably mediated by apoptosis (Allan et al., 
1992). Spontaneous degeneration of spermatogenic cells has also been reported to occur 
in humans (Johnson et al., 1984) with affected cells displaying characteristics of testicular 
cells undergoing apoptosis (Miething, 1992; Tapanainen et al., 1993; Brinkworth et al., 
1995). Miething (1992) reported that degenerating prespermatogonial germ cells in 
immature golden hamsters showed increased nuclear and cytoplasm staining intensity 
characteristic of decondensation. Here the nucleus degenerated into multiple fragments 
with formation of apoptotic bodies (with or without chromatin) which were eventually 
phagocytosed by Sertoli cells. In another study, Gorczyca et al. (1993) used human sperm 
to analyze DNA strand brakes by labeling 3'-OH with dUTP. A correlation of r=0.87 was 
observed between those cells with DNA strand brakes and cells that expressed high 
sensitivity to DNA denaturation, with the characteristics of this latter group resembling 
those of apoptotic somatic cells. 






17 
Effect of Gossvpol on Reproduction 

Gossypol is a toxic polyphenols pigment produced in the pigment glands of roots, 
leaves, stems and seeds of the cotton plant (Berardi and Goldbleatt, 1969). Gossypol has 
been related to reproductive problems in several species including humans, rodents and 
cattle (Liu, 1957; Liu and Segal, 1985, Jimenez et al., 1989; Randel et al., 1992; Chase et 
al., 1994; Chenoweth et al., 1994). It has been reported that gossypol increases free 
radical production (de Peyser et al., 1984; Barhoumi and Burghardt, 1996), suggesting 
that free radicals play an important role in gossypol toxicity. Male effects have been 
discussed previously. 

In female mice, when gossypol acetic acid was administered orally in doses of 40 
or 80 mg/kg body weight/d, pregnancy rates decreased from 90% in controls to 60 or 10% 
in gossypol treated animals respectively (Hahn et al., 1981). In another study, Lagerlof 
and Tone (1985), reported a decrease in pregnancy rates (89% in controls vs 60% in 
treated group) in female rats treated orally with gossypol acetic acid (20 mg/kg body 
weight/d). It has also been reported that gossypol decreased the ability of murine 
embryos to develop in vitro. In this study (Lin et al., 1989), development scores of 
embryos were significantly decreased (P<0.01) after 72 h of culture from 2.6 in control to 
1 .8 in embryos treated with 5.3 ng of gossypol compared with controls. Similar results 
were reported in bovine embryos by Zirkle et al. (1988). Here, embryos were cultured 
with 0, 1, 5, 10, or 30 /ug of gossypol, resulting in lower final development scores in 
treated than in control embryos (4.1, 3.3, 2.5, 0.4 or 0, respectively; PO.05). 



18 
Characteristics of Apoptosis or Programmed Cell Death 

Morphologically, apoptosis is characterized as occurring in a relatively small 
proportion of cells. In general, cells exhibiting apoptosis undergo shrinkage and 
condensation of chromatin and cytoplasm with chromatin fragmentation (Brinkworth et 
al., 1995). Eventually, the cell disrupts into randomly distributed organelles, bounded by 
membranes that are termed "apoptotic bodies." After apoptotic bodies are formed, 
affected cells are generally phagocytosed by macrophages. With light microscopy, 
apoptotic liver cells are seen to be surrounded by an abnormal halo and normal tissue, and 
exhibit condensed basophilic chromatin and eosinophilic cytoplasm. Apoptotic bodies 
may or may not contain chromatin, and affected cells are phagocytosed by macrophages 
or epithelial cells (Alison and Sarraf, 1992). Microvilli and junctions of plasma 
membranes are lost. The chromatin condenses and nuclear crescents are pushed to the 
periphery of the nucleus resulting in obscure nuclear pores and frequent blebbing of the 
nuclear membrane. Subsequently, nuclear fragmentation occurs and the nucleolus 
progressively disintegrates into osmiophilic granules. Integrity of the organelles is 
maintained, and mitochondria are still active until late stages of apoptosis. Brinkworth et 
al. (1995) suggested that testicular apoptosis may take a form characterized by cellular 
swelling and the appearance of decondensed, homogenous chromatin. In apoptosis, DNA 
is cleaved in an incremental manner of multiples of 180-200 base pairs associated with 
each nucleosome, giving a characteristic ladder pattern on gel electrophoresis. In 
apoptotic immature rat thymocytes, disruption of double-stranded DNA leads to the 



19 
formation of oligonucleosomes resulting in characteristic chromatin condensation 
patterns (Wyllie et al., 1980; Cohen and Duke, 1984; Wyllie et al., 1986). However, 
absence of this DNA ladder pattern has also been reported in cells undergoing apoptosis 
(Barbieri et al., 1992; Cohen et al, 1992), leading to some disagreement as to its 
diagnostic value as a marker for apoptosis. 

Characteristics of Necrosis 

In necrosis, cell death occurs in response to a wide variety of harmful conditions 
and toxic substances. Necrosis affects groups of continuous cells and inflammatory 
reaction usually develops in the adjacent viable tissues in response to the related cellular 
debris. Necrosis is characterized by swelling of the cytoplasm and organelles which leads 
to organelle dissolution and rupture of plasma membrane. These allow the cellular 
contents to leak out into the extracellular space, damaging neighboring cells, and the 
DNA is randomly cleaved by lysosomal deoxyribonuclease resulting in DNA fragments 
of different sizes (Wyllie, 1981; Allan and Harmon, 1986). 

Genes Reported to Play a Role in the Regulation of Apoptosis 

Apoptosis is a gene-regulated process. Apoptosis is observed in different cells 
and organs such as the developing embryo, immune system, ovary, testes, and others. In 
order to study the genes involved in the regulation of programmed cell death, the 
nematode C elegans has been used (Ellis et al., 1991). In C elegans, several genes have 
been mapped, and a series of genes have been described at different stages of apoptosis 
(Hale et al., 1996). In this nematode, Egl-1, ces-1, and ces-2 genes have been described 



20 
as genes that specify cell death (Ellis et al., 1991 ; Hale et al., 1996). In contrast ced-3, 
ced-4, ced-8, and ced-1 are genes that stimulate programmed cell death or apoptosis (Hale 
et al., 1996). Ced-9 is the gene that inhibits or suppresses programmed cell death in the 
C elegans (Hale et al., 1996). In this nematode, genes involved in the phagocytosis of 
dead cells or apoptotic bodies are ced-1, ced-6, ced-7, ced-2, ced-5, and ced-10. Nuc-1 is 
the gene that regulates digestion of dead cells (Hale et al., 1996). The C. elegans genes 
are important for the study of mammalian apoptosis since there is homology between 
mammalian genes involved in apoptosis and the genes regulating apoptosis in this 
nematode (Hale et al., 1996). 

In different species, including mammals, different genes have been described as 
programmed cell death inducers. Tumor suppressor p53, Apo 1 (FAS mediated 
pathway), nedd2, and ICE are genes that induce apoptosis in mammals (Donehower et al., 
1992; Hollstein et al, 1991; Kumar et al., 1992; Kumar et al., 1994; Nagata and Golstein, 
1995; Yuan et al., 1993). In the ced-9 family, a large groups of genes have been 
described that inhibit or suppress programmed cell death,. In humans, Bcl-2, Bax, Bcl-x, 
Mcl-1, and Bak-2 are included in the ced-9 gene family (Boise et al., 1993; Chittenden et 
al., 1995; Kozopas et al., 1993; Oltvai et al., 1993; Tsujimoto and Croce, 1986). In 
murines, ced-9 related genes, homologous to human genes have been described as well as 
2 murine specific genes (Al and Bad) (Lin et al., 1993; Yang et al., 1995). Also, viral 
genes like Epstein-Barr virus (BHRF1), African swine fever virus (LMW5-HL), and 
herpes virus Samiri (ORF16) have been described to induce apoptosis (Pearson et al., 
1987; Neilan et al., 1993; Smith, 1995). 



21 
Detection of Apoptosis in Testicular Tissue 

The determination of apoptosis in testicular tissue has been made with use of 
several methods including histomorphology of chromatin fragments, immuno- 
histochemical in-situ end-labeling of fragmented DNA, and radioactive DNA- 
fragmentation (Allan et al., 1992; Tapanainen et al., 1993; Troiano et al., 1994). 
However, quantification of apoptosis has been difficult with the above methods. In a 
more recent report (Hingst and Blottner, 1995), testicular apoptosis in sexually active 
guinea pigs, roe deer, and bulls was quantified in units per mg of testis (U/mg) using an 
ELISA test (7.08 ± 1.95, 16.32 ± 3.45, and 29 ± 7.1 U/mg testis, respectively). In another 
study, Blottner et al. (1995), using the same ELISA procedure, reported testicular 
apoptosis in the brown hare during the testicular proliferation phase of 14.16 ± 2.12 U/mg 
testis. Another procedure is the in-situ "Tunel" staining method which labels newly 
formed 3'-OH DNA ends of fragmented DNA in-situ, making possible the identification 
of specific cell types undergoing apoptosis (Li et al., 1995; Richburg and Boekelheide, 
1996; Hikimetal., 1997). 

Apoptosis in Testicular Tissue 

Although apoptosis may be particularly relevant as a putative cause of 
spermatogenic dysfunction in the male, relatively little is known concerning the causes or 
mechanisms of apoptotic processes in the male gonad, or their role in male infertility. 
Testicular apoptosis may be induced by a variety of agents. In the rat testis, increased 
temperature as a result of cryptorchidism resulted in increased numbers of apoptotic germ 



22 
cells (Shikone et al., 1994). Hormonal events influence apoptosis. Blottner et al. (1995) 
reported that apoptosis in testes of roe deer was significantly higher during the 
nonbreeding season than during the breeding season. Interestingly, Haigh et al. (1984) 
observed an increase in diadem/crater defects of sperm in wapiti, also during the 
transitional period. Apoptosis in testicular tissue has also been reported after withdrawal 
of pituitary gonadotropins and testosterone. In the rat, testicular apoptosis increased both 
after hypophysectomy and following administration of a GnRH antagonist (Tapanainen et 
al., 1993). However, when hypophysectomized rats were treated with FSH-CTP, hCG, or 
testosterone, hypophysectomy-induced apoptosis was less evident (16, 49, and 25% 
respectively of levels in non-treated, hypophysectomized rats) (Tapanainen et al., 1993). 
Troiano et al., (1994) reported that apoptosis of haploid germ cells in the adult rat 
increased after testosterone withdrawal. Brinkworth et al. (1995) treated adult male rats 
with a GnRH antagonist or with methoxyacetic acid, a highly toxic agent for rat 
pachytene spermatocytes. The GnRH antagonist decreased gonadotropin secretion, as 
expected, whereas methoxyacetic acid did not cause significant hormonal changes. Both 
treatments induced the characteristic DNA ladder pattern of apoptosis, as determined by 
electrophoresis, and increased numbers of degenerating apoptotic germ cells, although 
some differences occurred in the types of spermatogenic cells affected. 



23 
Germ Cell Types Involved in Apoptosis 

All germ cells are capable of undergoing apoptosis. However, some types appear 
to be more susceptible than others, or at least they have been more commonly reported to 
undergo apoptosis. 

In the golden hamster, prespermatogonial cells have been reported to undergo 
apoptosis between 14 days post conception and 13 days post partum (Miething, 1992). In 
the normal rat testis, type A 2 A 3 , and A 4 spermatogonia have been reported to undergo 
spontaneous apoptosis (Allan et al., 1992). Shikone et al. (1994), observed that 
experimentally induced cryptorchidism in 22 d old Sprague-Dawly rats resulted in 
apoptosis of primary spermatocytes. However, when adult Sprague-Dawly rats were 
treated with saline, 1 12.5 /u.g of GnRH antagonist, or 650 mg of methoxy acetic acid daily 
for 14 days, A type spermatogonia, as well as leptotene, zygotene, early pachytene, 
secondary spermatocytes, and spermatids all underwent apoptosis. In GnRH antagonist 
and methoxyacetic acid treated rats, the types of germ cells undergoing apoptosis were 
pachytene spermatocytes (all stages), preleptotene stage VII and VIII spermatocytes, and 
stage XII-XIII spermatocytes (Brinkworth et al, 1995). Richburg and Boekelheide 
(1996) treated 28 day old Fischer male rats with mono-(2-ethylhexyl) phthalate, resulting 
in apoptosis of spermatogonia. In a different study, Cai et al. (1997) treated Sprague- 
Dawley male rats with 70 mg/kg of cyclophospamide, inducing apoptosis in all germ 
cells. However, apoptosis was most evident in spermatogonia and in stage I-IV and XI- 
XIV spermatocytes. 



24 
Heat Shock Proteins 

A variety of cells synthesize heat shock proteins (Hsp) in response to stressors 
such as hyperthermia (Malayer et al., 1988; Putney et al., 1988; Gutierrez and Guerriero, 
1991 ; Harris et al, 1991 ; Edwards et al., 1997). Heat shock proteins are important for 
protein oligomerization, folding, translocation, secretion, and recognition of malfolding 
proteins, as well as protecting ribosomal RNA (Riabowol et al., 1988; Duncan and 
Hershey, 1989; Henry and Kola, 1991; Nover and Scharf, 1991). 

In the bovine, Hsp70 has been reported to be induced in response to heat shock in 
endometrial tissue, bovine embryos, lymphocytes, skeletal muscle, brain, kidney, liver, 
lungs, spleen, testes, and others (Malayer et al., 1988; Putney et al., 1988; Gutierrez and 
Guerriero, 1995; Edwards et al., 1997). Levels of Hsp70 in bovine tissues are shown in 
Table 2-1. Also, levels of Hsp70 in lymphocytes cultured at 42° C for 1 h were 5.0 ± 
0.36, 4.8 ± 0.36, and 4.2 ± 0.39 Mg/mg of protein for Angus, Brahman, and Senepol 
heifers, respectively (Kamwanja et al., 1994). 

Heat Shock Proteins in Testicular Tissue 

Spermatozoa possess highly condensed chromatin and are thus unable to undergo 
transcription. However, O'Brien (1987) identified Hsp and other proteins as being 
synthesized in mouse leptotene/zygotene spermatocytes, pachytene spermatocytes, and 
round spermatids. Allen et al., (1988) reported the presence of a 70 Kda protein related to 
Hsp 70, a novel protein synthesized in association with germ cell differentiation. Heat 
shock protein 70 is evident in small amounts in unstressed preleptotene and 






25 



Table 2-1. Concentration of Hsp70 in bovine tissues as determined by ELISA. 



Hsp70 Concentration (ng/,ug) ± SD 



Tissue 



Gutierrez and Guerriero, 1991 Gutierrez and Guerriero, 1995 

2.1 ±0.20 
4.1 ±0.30 
3.6 ±0.30 

3.6 ±0.70 
2.9 ±0.50 

8.7 ±0.40 
2.4 ±030 
2.6 ±0.20 



Brain 


1.9±0.15 


Heart 


3.5 ±0.06 


Kidney 


5.5 ±0.32 


Liver 


4.7 ±0.20 


Lung 


3.4 ±0.63 


Skeletal Muscle 


9.1 ±0.71 


Spleen 


2.6 ±0.60 


Testes 


1.8 ±0.20 



26 
leptotene-zygotene spermatocytes, but in larger amounts in pachytene spermatocytes 
(where most synthesis occurs) and in round spermatids. Heat shock protein 72 has been 
induced in male mouse germ cells by elevated temperatures (Zakeri et al., 1990). Miller 
et al. (1992) have identified a number of heat shock proteins in human sperm. 

Scrotal Insulation Model 

The scrotal insulation model has been widely used to study adverse effects upon 
spermatogenesis in male ruminants, with its effects on aspects of bull spermatogenesis 
being particularly well characterized. These effects include reduced spermatogenesis, 
consistent temporal patterns of occurrence of different sperm abnormalities, and adverse 
effects on sperm freezability. For example, Austin et al. (1961) compared scrotal 
insulation times of either 24 h (n=4) or 72 h (n=4) for mature bulls. Both treatments 
caused a decline in both live and normal sperm to 55% of control levels by 2 weeks 
following treatment. Mean sperm concentration also declined following both treatments 
to 60% of control levels at 6 weeks. Scrotal insulation for 6-1 1 weeks in 10 healthy men, 
ranging from 1 9-43 years old, resulted in a depression of spermatogenesis to a mean 
14.1% of pre-treatment levels at the 6th week of treatment (Robinson and Rock, 1967). 

Ross and Entwistle (1979), using five normal bulls, applied scrotal insulation for 
either 10 or 20 h. Sperm concentration in ejaculates declined in the 10 h group only, 
decreasing from weeks 7 to 1 1 post treatment. These workers also described testicular 
cell degeneration (and types of cells involved) following scrotal insulation by injection of 
3 H-thymidine into the spermatic artery. Testicular sections were taken for histology after 



27 
1 .25-2 h or 13.6-14.5 d of 3 H-thymidine infusion and scrotal insulation (10 or 20 h). In 
both treatments, the number of B-type spermatogonia and young spermatocytes declined 
in the sections taken around d 14 when compared with the sections taken 1-2 h following 
3 H-thymidine infusion. 

The effect of scrotal insulation on cryopreserved bovine sperm has also been 
documented. Vogler et al. (1991) reported that scrotal insulation for 48 h in six young 
bulls lowered sperm viability in semen collected from d 12-39 after insulation. Semen 
collected at d 3-9 that was frozen/thawed and incubated for 3 h at 37° C exhibited lower 
spermatozoal viability than semen collected prior to treatment. 

Bull sperm abnormalities following 48 h of scrotal insulation were described by 
Vogler et al. (1993). Significant increases in abnormal sperm became apparent 12 d after 
initiation of treatment (47.5 ± 27.4%), peaking at d 18 (86.3 ± 9.8 %). Lowest sperm 
motility occurred 15 d after treatment (42 ± 9.8 %). Different types of sperm 
abnormalities predominated in a sequential manner beginning with tailless sperm at d 12- 
15, "diadem" sperm defects at d 18, nuclear vacuoles and pyriforme heads at d 21, 
knobbed acrosomes at d 27, and "Dag" defects at d 30. 

Scrotal skin temperature has been reported to increase as a result of scrotal 
insulation. Austin et al. (1961) reported a skin temperature of 33.3° C immediately after 
initiation of insulation, with mean scrotal skin temperatures as a result of insulation being 
33.9°, 35.4°, 36.1 °, and 34.7° at 8 a.m., 12 noon, 4 p.m., and 10 p.m. respectively. Ross 
and Entwistle (1979) reported a maximum temperature in the scrotal pouch of 35.0 ± 0.5° 
C, temperature during the period of scrotal insulation. Wildeus and Entwistle (1983a) 



28 
applied scrotal insulation for 48 h to bulls reaching scrotal temperatures ranging between 
32° to 34° C in controls vs 35.5° to 38° in scrotal insulated bulls, with a mean 
temperature difference of 4.05° C between scrotal insulated and controls. Sidibe et al. 
(1993) reported a mean testicular skin of 31.6° C before scrotal insulation and mean 
temperatures of 34.8°, 35.5°, 35.8°, 36°, and 35.8° C inside the scrotal insulator at 2, 24, 
48, 72, and 96 h after initiation of scrotal insulation, with a scrotal temperature increase 
of 3.2° to 4.4° C. Vogler et al. (1993) reported a mean testicular surface temperature of 
34.8° C as a result of scrotal insulation, with the temperature of the testicular surface 
ranging from 33.3° to 36.4° C. 

Overall these reports indicate that scrotal insulation caused an increase in scrotal 
skin temperature in the bull, and that this increase in scrotal skin temperature is enough to 
disrupt spermatogenesis in the bull. 

Effect of E. coli Endotoxin Infusion 

This endotoxin is a lipopolysaccharide derived from E. coH 055 :B5, which has 
been used in horses to study peritoneal macrophages and to induce mastitis in the bovine 
(Morris et al., 1992). In the horse, infusion of E. coli endotoxin causes pyrexia and an 
increase in plasma concentration of TNF, IL-6, 6-keto PGFla, thromboxane, and other 
compounds. Escherichia coli endotoxin increased IL-6 when administered to neonatal 
foals (Robinson et al., 1993), and when 1000 or 30 ng/kg of endotoxin were administered 
to the horse, IL-6 increased after 3 h of infusion from 171 ± 10.2 U/ml to 10128 ± 4096 
and 1555 ± 1326 U/ml, respectively; TNF was detectable in blood and pyrexia occurred 



29 
(MacKay and Lester, 1992). In another experiment (Morris et al., 1992), horses infused 
with 30 ng/kg of endotoxin showed pyrexia, leukopenia after 1 h, leukocytosis after 8 h, 
and an increase in IL-6 between 1 .5 and 5 h after infusion, peaking between 3-4 h. 
Endotoxin also caused local increases of IL-6, TNF, and inflammatory signs when 
infused intra-articularly or into the mammary gland (Hawkins et al., 1993; Shuster et al., 
1993). 

Effect of Elevated Temperatures on Testosterone Levels 

Lowered hormonal levels are associated with increased spermatogenic apoptosis 
in several species such as deer and rat (Tapanainen et al., 1993; Troiano et al., 1994; 
Brinkworth et al., 1995; Blottner et al., 1996), and this finding is relevant for bulls in 
tropical or subtropical regions of the world. Here, heat stress may lower levels of 
pituitary gonadotropins and testicular androgens (Rhynes and Ewing, 1973; Minton et al., 
1981; Sidibe et al., 1992), increasing apoptosis levels in germ cells. Peripheral 
testosterone levels in bulls have shown variable effects as a result of environmental heat 
stress. This is possibly because the testicles of bulls, in common with a number of 
domestic species, have the ability to preserve its temperature (thermoregulation) through 
complex physical and physiological mechanisms. Rhynes and Ewing (1973) reported that 
plasma testosterone levels of 8 mature (20-26 months old) Hereford bulls decreased to 
43% of control values during exposure to elevated temperature (35.5 ± 1° C) for 7 weeks 
in environmental chambers following a 7 week control period (21.1 ± 1° C). In contrast, 
Minton et el. (1981), reported that serum testosterone was not reduced in bulls that were 



30 
exposed to 34° C in environmental chambers for 15 days. Peripheral testosterone 
concentrations have decreased after local heating of the testis. For example, Sidibe et al. 
(1992), using 3 year old bulls, found that serum testosterone levels decreased by 25% 
following 96 h of scrotal insulation, reaching lowest levels at 6 weeks following 
treatment. Setchell et al. (1991) reported that exposure of ram testes to 42° C for 45 
minutes resulted in lowered peripheral plasma levels of testosterone and lowered 
testosterone levels in rete testis fluid. However, plasma testosterone levels in blood 
collected from the spermatic vein did not differ whereas testicular blood flow was lower 
in heat treated testicles than in controls. Similar results have been observed in rats. Galil 
and Setchell (1987) reported lower testosterone concentrations in peripheral blood from 
testis-heated (41.5-43° C) rats following stimulation with hCG 21 d post heating. In 
contrast, the testosterone concentration in blood collected from testicular veins after hCG 
stimulation was higher in heat-treated rats than controls. These results suggested that 
changes in testosterone concentration resulting from elevated temperatures are caused by 
changes in testicular blood flow, and changes are not necessarily due to decreased Leydig 
cell capability to synthesize testosterone. 

Testosterone concentrations also decline with testicular involution during the non- 
breeding season in seasonal breeders. Finnish Landrace and Suffolk rams have lower 
testosterone concentrations in the non-breeding season than in the breeding season 
(Schanbacher and Lunstra, 1976). Similar results were reported in roe deer by Blortner et 
al. (1995), with testosterone concentrations being highest during the breeding period. 



31 
Effect of Elevated Temperatures on Gonadotropins 

Pituitary gonadotropins can also be adversely affected by elevated temperatures. 
Minton et al. (1981) exposed eight mature bulls to 34° C using environmental chambers 
to evaluate the effects of elevated ambient temperature on serum LH before and after 
challenge with GnRH. Bulls were exposed during a 3 week period at 22° C. After this 
adjustment period bulls were separated into control (22 ± 1° C) or heat stress (34 ± 1° C) 
groups for a 15 d treatment period. Blood samples were collected on days -2, 6, and 15 of 
treatment at 30 minute intervals for a 12 h period, after which the bulls were parenterally 
administered 200 ug GnRH. Blood samples were subsequently taken every 1 5 minutes 
for 1 h and then every 30 min for 5 h. Non-stimulated LH levels were reduced on days 6 
and 15 in heat-stressed bulls compared to controls. In addition, LH secretory peaks were 
reduced in heat-stressed bulls compared with controls on days 6 and 15. However, LH 
levels in response to GnRH challenge were not reduced by heat stress. In contrast, when 
testes of rams were heated by immersing the scrotum in a water bath (45° C) for 30 or 45 
minutes, plasma LH and FSH concentrations did not differ in heat treated rams as 
compared to controls (Setchell et al., 1991). This differed from a previous study by Galil 
and Setchell (1987) where exposure of rat testes to 43° C for 30 minutes caused increases 
in plasma LH and FSH concentrations from d 7 to d 42 following treatment. 



32 
Breed Differences in Susceptibility to Elevated Temperatures 

Breed differences in susceptibility to elevated temperatures have been reported 
between Bos taurus and Bos indicus genotypes. For example, Johnston et al. (1 963) 
exposed purebred Holstein Friesian, Brown Swiss, and Red Sindhi crossbred bulls to a 
maximum of 104° F and 54% relative humidity and a minimum of 82° F and 72% 
relative humidity for an 8-h period every day for 7 days using environmental chambers. 
Crossbred bulls (Red Sindhi x Holstein and Red Sindhi x Brown Swiss) had lower 
(PO.01) rectal temperature (100°± 0.12 and 102.8°± 0.27; 101.5°± 0.12 and 103°± 0.32 
F, respectively) when air temperature was 82° or 104° F than purebred bulls (Holstein, 
102.6°± 0.28 and 105.2°± 0.35 F; Brown Swiss, 103.6°± 0.30 and 104° ± 0.34 F). 
Respiration rates were also lower (PO.05) in crossbred (56 ±3.9 and 60 ± 5.5 
counts/min, respectively) than in purebred bulls (80 ±3.2 and 75 ± 6.4 counts/min, 
respectively). Results showed that elevated temperatures and high humidity lowered 
semen quality, and that these deleterious effects were greater in purebreds than in 
crossbred bulls. 

In Florida, Fields et al. (1979), compared semen traits in yearling Hereford, 
Angus, Santa Gertrudis, and Brahman bulls for semen traits. In this study, Herefords 
from Montana showed lower semen quality than Brahman, Angus, Hereford from Florida, 
and Santa Gertrudis. In addition, breeds with better ability for thermoregulation such as 
Brahman are also more resistant to the effect of heat stress than more thermosensitive 
breeds such as the Scottish Highland (Seif et al., 1979). Another example of breed 



33 
differences in response to elevated temperatures was observed in Australia, where Turner 
(1982) measured rectal temperatures in two B. taurus (Hereford x Shorthorn) and four B. 
indicus x B. taurus lines. In this study the average rectal temperature was 39.8 °C, being 
0.5 ° C higher in B. taurus than in B. indicus x B. taurus crosses. 

Related differences in response to elevated temperatures have also been reported 
in Bos taurus and Bos indicus bulls in the tropics. For example, Kumi-Diaka et al. 
(1981), reported higher sperm abnormalities, lower percentage live-sperm, and lower 
sperm concentration during the hot periods in Bos taurus than in Bos indicus bulls. 

The Senepol is a tropically adapted B. taurus breed developed in St. Croix US 
Virgin Islands during the early 1900s. Senepol originated from crosses between the 
N'Dama and Red Poll breeds (Hupp, 1981). The Romosinuano is another tropically 
adapted B. taurus breed. Romosinuano is a criollo breed native to Colombia (Rouse, 
1977). Bos taurus tropically adapted breeds have been reported to be more resistant to 
heat stress than temperate B. taurus such as the Angus (Chase et al., 1993; Hammond et 
al., 1996). For example, when temperate B. taurus (Angus) bulls were compared with 
tropically adapted B. taurus (Senepol) during the Florida summer, Senepol bulls had 0.5° 
C lower (P<0.003) rectal temperature than Angus bulls; semen quality tended to decrease 
in Angus but not in Senepol bulls (Chase et al., 1993). When Senepol bulls were 
compared with Holstein bulls (temperate B. taurus ) in the semi-arid environmental 
conditions of St. Croix, Virgin Islands, Wildeus and Hammond (1993) reported that 
Senepol bulls had lower (PO.01) rectal temperatures than Holstein bulls (39.3 vs 40.0° 
C), and packed cell volume was higher (PO.01) in Senepol than in Holstein bulls (41.1 



34 
vs 35.2%, respectively). Also, Senepol bulls showed overall higher spermatozoa 
concentration with lower percentage of sperm abnormalities than Holstein bulls (P<0.05). 

In another study illustrating physiological differences between genotypes, 
Kamwanja et al. (1994), cultured lymphocytes from Angus (temperate Bos taurus), 
Brahman (tropically adapted Bos indicus), and Senepol (tropically adapted Bos taurus ) 
heifers (n=12 per breed) to 45° C for 3 h. In this experiment cell death was affected by a 
breed by temperature interaction (P<.01), with cell viability being lower for Angus than 
for Brahman or Senepol, indicating that lymphocytes from tropically adapted breeds 
(Brahman and Senepol) were more resistant to heat shock than lymphocytes from Angus. 
In addition, Hammond et al. (1996) evaluated heat tolerance among temperate B, taurus 
(Angus and Hereford), B. indicus (Brahman), and tropically adapted B, taurus (Senepol 
and Romosinuano) heifers. Here, rectal temperature was lower (P<0.001) in Brahman, 
Senepol, and Romosinuano than in Angus heifers (39.6°, 39.2°, and 39.5° vs 40.4° C, 
respectively), and respiration rates were faster (P<0.05) in Angus and Hereford than in 
Brahman, Senepol, and Romosinuano (69 and 64 vs 36, 57, and 55 respirations per min, 
respectively). These results suggest that spermatogenesis, a temperature dependent 
process, could be less disrupted in tropically adapted than in temperate breeds under heat 
stress conditions. 

Hammond et al. (1998) investigated heat tolerance in Tuli x Angus, Senepol x 
Angus, and Brahman x Angus heifers under environmental conditions in central Florida. 
In trial 1, 38 Brahman, 21 Senepol, 19 Brahman x Angus, 20 Senepol x Angus, and 20 
Tuli x Angus heifers were used. Rectal temperature (log, ) on the hottest day did not 



35 
differ between Brahman and Brahman x Angus (0.39 ± 0.01 1 and 0.37 ± 0.016, 
respectively) or between Tuli x Angus and Brahman x Angus (0.35 ± 0.015 and 0.35 ± 
0.015, respecively). Rectal temperature (log 10 ) was lower in Senepol x Angus than in 
Senepol and Tuli x Angus (0.30 ± 0.015 vs 0.35 ± 0.015 and 0.35 ± 0.015 respectively; 
PO.05), and respiration rate and blood packed-cell volume were lower (PO.05) in 
Brahman than in Brahman x Angus. In trial 2, 13 Angus, 35 Brahman, 30 Senepol, 23 
Brahman x Angus, 17 Senepol x Angus, and 28 Tuli x Angus heifer were used. On the 
hottest day of trial 2, rectal temperature (log I0 ) and respiration rate were higher (P<0.05) 
in Angus (0.59 ± 0.017 and 74 ± 2.7, respectively) than Brahman (0.47 ± 0.01 and 39 ± 
1.6, respectively), Tuli x Angus (0.47 ± 0.01 1 and 60 ±1.8, respectively), Senepol x 
Angus (0.43 ± 0.014 and 55 ± 2.4, respectively), and Tuli x Angus (0.50 ± 0.012 and 48 ± 
2.0, respectively). Also, respiration rate was higher (PO.05) in Brahman x Angus than in 
Brahman. This work indicates that heat tolerance in female crosses between tropically 
adapted Brahman, Senepol, and Tuli with temperate Angus is similar to that of tropically 
adapted Senepol or Brahman. 

As elevated temperatures disrupt spermatogenesis (Johnston and Branton, 1953; 
Casady et al., 1953; Johnston et al., 1963; Vogler et al., 1993), it is possible to speculate 
that non-heat-tolerant breeds such as temperate B. taurus (Angus) are more susceptible to 
suffer more spermatogenic damage as a result of elevated temperatures than heat-tolerant 
breeds such as B. indicus (Brahman) or tropically adapted B, taurus (Senepol or 
Romosinuano). Also, if non-heat-tolerant breeds undergo more spermatogenic damage 
than heat-tolerant breeds, it is possible that spermatogenic apoptosis could be higher in 



36 
temperate B. taurus (non-heat-tolerant) than in tropically adapted or heat-tolerant B. 
indicus, B. taurus , and B. indicus x B. taurus crossbred breeds. No references were, 
however, found which addressed this specific question. 



CHAPTER 3 
ATTEMPTS TO INDUCE AND ASSESS SPERMATOGENIC APOPTOSIS IN BULLS 



Introduction 

Male infertility has been associated with many stressors such as elevated 
temperatures, toxic agents, and disease processes (Vogler et al., 1993; Richburg and 
Boekelheide, 1996; Burgess and Chenoweth, 1975). In the bull, environmental heat stress 
is probably the most ubiquitous. In the southern subtropical US, cattle fertility is 
significantly lower than in more temperate zones (Chenoweth, 1 994), with heat stress 
being a great contributor to the problem (Badinga et al., 1985; Weller and Ron, 1992). 
Heat stress is also implicated as a cause of decreased spermatozoal quality in ejaculates of 
bulls during the summer months (Fields et al., 1979). Spermatogenic damage in several 
species follows apoptosis, an active process of gene-directed cellular self-destruction 
(Kerr and Harmon, 1991). Increased spermatogenic apoptosis has been described in 
several species as a result of elevated temperature, spermatoxicity, hormonal withdrawal 
and seasonal testicular involution (Shikone et al., 1994; Brinkworth et al., 1995; Blottner 
et al., 1996; Hikim et al., 1997). 

Although spermatogenic apoptosis has been associated with spermatogenic 
dysfunction in the male, relatively little is known concerning its causes, mechanisms, and 
role in male infertility. In many species, the spontaneous death of testicular germ cells 

37 



38 
appears to be common (Roosen-Runge, 1973; Allan et al., 1987). Normally, sperm output 
is often lower than expected in mice, rats, and humans when compared with the 
theoretical number that should be obtained from spermatogonial precursors (Oakberg, 
1956; Russel and Clermont, 1977; Johnson et al., 1984; Kerr, 1992). The contribution of 
apoptosis to sperm reduction is unknown and may only be hypothesized at present, even 
though apoptosis is often observed in spermatogenic tissue. Kerr (1992) described 
spermatogonial depletion as a cause of testicular germ-cell loss in rats, an effect 
presumably mediated by apoptosis (Allan et al., 1992). Spontaneous degeneration of 
spermatogenic cells has also been reported to occur in humans (Johnson et al., 1984) with 
affected cells displaying characteristics of testicular cells undergoing apoptosis (Miething, 
1992; Tapanainen et al., 1993; Brinkworth et al., 1995). Miething (1992) reported that 
degenerating prespermatogonial germ cells in immature golden hamsters showed 
increased nuclear and cytoplasm staining intensity characteristic of decondensation. He 
reported that the nucleus degenerated into multiple fragments with formation of apoptotic 
bodies (with or without chromatin) which were eventually phagocytosed by Sertoli cells. 

In the present study, two experiments were designed to test the hypothesis that 
spermatogenic stress, caused by elevation of either body temperature by E. coli endotoxin, 
or testicular temperature by scrotal insulation, will increase both spermatogenic 
dysfunction and spermatogenic apoptosis in bulls. Additional objectives were to test 
breed differences in susceptibility to spermatogenic apoptosis as a result of scrotal 
insulation, and to determine changes in levels of spermatogenic apoptosis with respect to 
time of assessment following elevated temperature. 



39 

Materials and Methods 

Materials 

R coli 055:B5 endotoxin, Merthiolate, butanol, hydrogen peroxide (30%), and 
methyl green were purchased from Sigma Chemical Co. (St. Louis, MO). Sodium 
chloride, sodium acetate, Permount, xylene, and triton X- 1 00 were purchased from Fisher 
Scientific (Fair Lawn, NJ). Dulbeco's phosphate-buffered saline (DPBS) was purchased 
from Life Technologies Inc. (Grand Island, NY). Cell death detection ELISA-kits were 
purchased from Boehringer Mannheim Co. (Indianapolis, IN), and the in-situ Tunel 
staining kits (ApopTag peroxidase S7101) were purchased from Oncor (Gaithesburg, 
MD). Ethanol was purchased from AAPER Alcohol and Chemical Co. (Shelbyville, KY), 
and acetic acid was purchased from Scientific Products (McGaw Park, IL). 
Experiment 1 

Eleven mature Angus bulls (averaging 3 yr of age and 700 kg of body weight) of 
normal reproductive status were acclimatized for 1 wk, at the College of Veterinary 
Medicine, University of Florida, Gainesville, Florida, where a breeding soundness 
evaluation (BSE) was performed (d -3). Bulls were distributed into three experimental 
groups: 1) control, sterile saline infusion (n=3), 2) E. cob 055:B5 endotoxin infusion (100 
ng/kg) (n=4), and 3) scrotal insulation (SI) for 48 h (n=4). All treatments started on day 
0, when clinical signs (rectal temperature, respiration, and heart rates) were measured at 
0, 15, 30, 60, 120, 240, and 360 min after infusion. Insulation of the scrotum was 



40 
achieved by placing the scrotum in a sack made from two layers of waterproof nylon 
taffeta filled with a 1 cm thick batting (Vogler et al., 1993). The jugular vein was used to 
infuse the endotoxin or saline by venipuncture. 

On d 8, semen was collected using a Lane IIIZ Pulsator (Lane Manufacturing, 
Denver, CO) in conjunction with a 3 ventral-electrode rectal probe. Immediately after 
semen collection, sperm motility was evaluated, and a smear stained with nigrosin-eosin 
(NE) (Society for Theriogenology, Hastings, NE) was prepared. A minimum of 200 
spermatozoa per slide were assessed for morphology using oil-immersion bright-field 
microscopy (x 1 000), and abnormal sperm were classified based on the region (head, 
acrosome, midpiece, or tail) where the lesion occurred (Chenoweth et al., 1994). Also, 
semen was fixed in buffered isotonic formal-saline (FBS) and examined for abnormalities 
using differential-phase (DIC) microscopy employing a Zeiss Axioscope microscope (x 
1000) (Chenoweth et al., 1994). 

Animals were sacrificed on d 10 using standard industry procedures at the 
University of Florida Meats Laboratory (Gainesville, FL) or at Central Packing Inc. 
(Center Hill, FL). The timing of sample collection (i.e. collection of testicular tissue on 
10 d after initiation of treatments) was based on estimation of optimal timing following 
the work of Ross and Entwistle (1979), who examined testicular sections taken after 1.25- 
2 h or 13.6-14.5 d of 3 H-thymidine infusion and scrotal insulation (10 or 20 h). In both 
treatments, the number of B-type spermatogonia and young spermatocytes declined in 
sections taken at 14 d when compared with sections taken 1-2 h following 3 H-thymidine 
infusion. Thus, affected spermatogenic cells should be detected between 1 and 14 d 



41 
following treatment, and in this experiment d 10 was chosen. In the present experiment, 
testes were collected into plastic bags and immediately transported on ice to the 
laboratory. The spermatic cord was trimmed, and the intact testis and tunic were 
weighed. The tunic was then removed and the tunic and the testis without tunic were 
weighed separately. Testicular circumference was measured, and testis length, depth, and 
width were obtained using calipers. Paired testicular volume was calculated as the sum 
of the volume of the right and the left testes. Testes were considered as paraboloids, and 
their volume was calculated using the equation v = m^h.. Where r = (width + depth) / 4 
and h = length (Fields et al., 1979; Chase, et al., 1997). Epididymides were removed, 
weighed, and mid-parenchymal sections were taken for in-situ Tunel staining, for 
spermatogenesis quantification, and for ELISA. Mid-parenchymal sections were fixed in 
Bouin's fixative for histological evaluation of spermatogenesis (andrological evaluation 
or cell association) and apoptosis. 

The cell association assessment (andrological evaluation) was done as described 
by Berndtson et al. (1987). Briefly, Sertoli cells, type A spermatogonia, pre-leptotene, 
pachytene primary spermatocytes, and stage 8 round spermatids were counted in 20 cross 
sections of seminiferous tubules at stage VIII. Only Sertoli cells whose nucleus contained 
a nucleolus were counted. The resulting counts (crude counts) were used to calculate 
Sertoli : germ cell ratios. Cells with histological characteristics of apoptosis (basophilic 
and condensed nucleus with smooth edges) were counted in the 20 tubules used for the 
cell association assessment. 



42 
An in-situ Tunel staining method using the ApopTag S7101 in 10% buffered 
formalin fixed tissue was used to detect apoptosis (Hikim, et al., 1995). This method is 
based on the specific binding of terminal deoxynucleotidil transferase (TdT) to the newly 
formed 3'-OH of cleaved DNA. Testicular tissues were fixed in 10% neutral buffered 
formalin for 24 h, paraffin-embedded and sectioned. Sections were de-paraffinized by 
two washes in xylene (slide was dipped in and out once per wash), two washes in 1 00% 
ethanol, one wash in 90% and 70% ethanol, and DPBS. Endogenous peroxidase was 
quenched by using 2% hydrogen peroxide in DPBS and then rinsing twice with DPBS. 
Slides were placed in equilibrium buffer and then in working strength TdT enzyme. The 
reaction was stopped by adding working-strength stop/wash buffer. Two drops of anti- 
digoxigenin-peroxidase were applied to slides, and the peroxidase was detected with 
diaminobenzidine. Negative controls were prepared by adding distilled water instead of 
TdT enzyme during the preparation of working-strength TdT. Sections were counter- 
stained with 0.5% methyl green (w:v in 0.1 M sodium acetate, pH 4), and slides were 
mounted using Permount. The number of apoptotic cells (stained positive) were counted 
in 10 seminiferous tubules. 

Apoptosis was quantified using a cell-death detection ELISA-kit, using aliquots of 
homogenized testicular parenchyma in DPBS (Hingst and Blottner, 1995). The assay is a 
quantitative sandwich-enzyme-immunoassay using monoclonal mouse antibodies against 
DNA and histones. The specificity of this ELISA permits determination of mono- and 
oligonucleosomes in the cytoplasmic fraction of cell ly sates. Testicular parenchyma (1 g 
in 2 ml DPBS) was minced, freeze/thawed (- 20° C/18° to 25° C) 3 times, and 



43 
homogenized for 1 min using a Polytron (Brinkman Instruments, Westbury, NY), 
followed by sonication for 1 min at 4° C and centrifugation at 25000 g for 30 min at 4°C. 
The supernatant was collected and stored in 100 ul aliquots at -20° C. Samples were run 
in triplicate. The microtiterplate-modules (MM) were coated with 100 ul of coating 
solution containing the anti-histone antibody, and cultured overnight at 4°C. Incubation 
buffer (200 ul) was pipetted into each well, wells were covered with plate cover foils, and 
incubated at room temperature (18° to 25° C) for 30 min to saturate the non-specific 
binding sites on the wall. After incubation, the incubation buffer was removed, and MM 
were washed 3 times with 300 ul washing solution. Then 100 ul of each sample solution 
containing 10 ug of testis equivalent/ 100 ul were added to the MM, covered, and 
incubated for 90 min at room temperature. For background determination, 100 ul of 
incubation buffer were pipetted into3 wells. Then, 100 ul of anti-DNA-peroxidase were 
added to the MM and incubated for 90 min at room temperature. After incubation, 
ABTS® (Boehringer) substrate was added to the wells, incubated at room temperature for 
10 min, and the absorbance read at 405 nm. The ELISA test was confirmed by running a 
standard curve using 0, 2.5, 5, 10, 20, 40, and 80 /ug of testis equivalent from 3 different 
bulls. 

Spermatogenesis was quantified by counting elongated spermatids to determine 
daily sperm production (DSP) and DSP/g (DSPG) of testicular parenchyma (Chenoweth 
et al., 1994). Briefly, 5 g of testicular parenchyma were thawed and finely minced. The 
sample was homogenized for approximately 1 min in 25 ml of working solution (0.9% 
NaCl, 0.05% Triton X-100, and 100 ppm Merthiolate diluted 1 :4 with distilled water). 



44 
Then, 175 ml of working solution were added and mixed for 1 min. The solution was 
allowed to settle for at least 1 h, and after the settling period, thoroughly mixed using a 
magnetic stirrer. Elongated spermatids were counted using a hemocytometer (4 fields), 
and the values were used to determine DSPG and DSP using the formula: 

DSPG = AX (B+Y) / (Time divisor)Y 
DSP = DSPG (0.99Z) 
Where X = hemocytometer count, Y = parenchyma sample weight, Z = testis parenchyma 
weight, A = hemocytometer constant, B = dilution factor, and time divisor (5.32 = time 
divisor for Bos taurus) (Amann et el., 1974). 
Experiment 2 

This experiment was conducted at the Subtropical Agricultural Research Station 
(28° 37' N latitude, 82° 22' W longitude), Brooksville, Florida. Bos taurus bulls (18 mo 
of age; Angus, An, n = 8; Senepol, Se, n = 6; Romosinuano, Ro, n = 4) were allotted to 
two experimental groups; 1) control (An, n = 4; Se, n = 2; Ro, n = 2) and 2) scrotal 
insulation (SI) for 48 h (An, n = 4; Se, n = 4; Ro, n = 2). Scrotal insulation began at d 0. 
Bulls were electro-ejaculated on d -6, d -4, and d 1, and semen was evaluated for motility 
and for morphology (NE and FBS) as described in experiment 1 . Semen samples were 
collected, placed on ice, and transported to the laboratory at the College of Veterinary 
Medicine (Gainesville, Florida). Samples were centrifuged for 30 min using a 
microcentrifuged, the supernatant was collected an kept frozen at - 20° C. Later, samples 
were shipped overnight on dry ice to the Department of Biomedical Sciences (Ontario 



45 
Veterinary College, University of Guelph, Ontario, Canada) for quantification of Hsp70 
in semen. Heat shock protein 70 was quantify using an ELISA test (unpublished data, 
Kamarundi and King, 1998). On d 1, rectal temperature and temperature inside the 
scrotal insulator was measured using a digital thermometer (HH 2 1 , Omega, Stamford, 
CT). At d 2, after 48 h of treatment, SI was removed from all animals, and 9 bulls were 
sacrificed within 2 h (control; An, n = 2; Se, n = 1 ; Ro, n = 1 ; and SI; An, n = 2; Se, n = 
2; Ro, n = 1). The remaining 9 bulls were sacrificed on d 4 (control; An, n = 2; Se, n = 1 ; 
Ro, n = 1 ; and SI; An, n = 2; Se, n = 2; Ro, n = 1). In a recent study, mono-(2-ethylhexyl) 
phthalate was shown to induce testicular apoptosis in the rat, with apoptosis increasing 
after 6 h of treatment (Richburg and Boekelheide, 1996). This suggested that detection of 
testicular apoptosis should be attempted earlier than in experiment 1 . As a result, 
detection of testicular apoptosis in experiment 2 was attempted either in tissue harvested 
within 2 h of removal of scrotal insulators (2 d) or within 2 days following this (4 d). 
After sacrifice, tissues were collected and processed as described in experiment 1 . 
Statistical Analysis 

Data were analyzed by least squares analysis of variance (ANOVA), using the 
GLM procedure of SAS (SAS, 1989, 1996). In experiment 1, the model included 
treatment and the error term was the residual. For clinical data, sperm motility, and FBS 
sperm morphology, treatment, bull within treatment, time, and the interaction of 
treatment by time were included in the model. The difference in sperm motility was 
calculated by deducting sperm motility at d 8 from sperm motility at d -3 (d -3 - d 8), and 



46 
the resulting values were analyzed. Bull within treatment was used as the error term for 
treatment, and the residual was used as the error term for the rest of model. In experiment 
2, the model included treatment, breed, treatment by breed, day, treatment by day, breed 
by day, treatment by breed by day, and the residual was used as the error term. Since no 
significant effects were observed, additional analysis including either treatment, day, and 
day by treatment, or treatment, breed, breed by treatment were conducted. The difference 
in sperm motility was calculated by deducting sperm motility at d 1 from the average 
sperm motility between d -6 and d -4, and the resulting values were analyzed. Data for 
spermatogenesis was also analyzed for breed, and in this analysis treatment was not 
included in the model. Data for rectal and scrotal temperatures were also analyzed 
following logarithmic transformations. All data are presented as least squares means ± 
SEM (LSMeans ± SEM). 

Results 

Experiment 1 

The effect of & coh endotoxin infusion (100 ng/kg) on (LSMeans ± SEM) 
respiration and heart rates as well as on rectal temperature is shown in Figure 3-1 . There 
was an effect of time of assessment on respiration rate and rectal temperature (P<0.05 
respectively), but not of treatment. Percent motility and FBS morphology values 
(LSMeans ± SEM) measured three days before the beginning of treatments (d -3) and 
eight days after beginning of treatment (d 8) are shown in Figure 3-2, Figure 3-3, and 
Table 3-1, respectively. Sperm motility and FBS sperm morphology (percentage of 



47 
normal spermatozoa, primary abnormalities, and secondary abnormalities) did not differ 
as a result of R cob endotoxin infusion or scrotal insulation (48 h) treatment or time 
when compared with control bulls. Least squares means (± SEM) for semen and 
testicular traits at d 8 in control, E cob endotoxin infused, or scrotal insulated (48 h) 
bulls are presented in Table 3-2. There was no effect of treatment on testicular and semen 
traits. Sertoli cells, germ cells, and ratio (LSMeans ± SEM) are shown in Table 3-3 and 
Figure 3-4. Sertoli cells, germ cells, and Sertoli:germ cell ratio did not differ among 
groups (control, endotoxin, or SI). Spermatogenic apoptotic cells (LSMeans ± SEM) 
determined by histology in control, endotoxin, or SI treated bulls are shown in Table 3-3 
and Figure 3-5. Spermatogenic apoptotic cells per seminiferous tubule determined by 
histology were not influenced by treatment (control, endotoxin, or SI). Results on 
apoptotic cells per seminiferous tubule (LSMeans ± SEM) determined by tunel stain are 
shown in Figure 3-6. Number of apoptotic cells (stained positive by tunel) per 
seminiferous tubule did not differ as a result of treatment. 

Confirmation of the ELISA test was done by a standard curve. A standard curve 
is shown in Figure 3-7. Results showed an effect of concentration (P< 0.05) on light 
absorbance at 405 nm. The ELISA test was able to detect differences between 
concentrations used (contrast vs 2.5, 5, 10, 20, 40, and 80 /*g, P< 0.05; 2.5 vs 5, 10, 20, 
40, and 80 /ug, P< 0.05; 5 vs 10, 20, 40, and 80 /zg, P< 0.05; 10 vs 20, 40, and 80 ^g, P< 
0.05). Least squares means (± SEM) for spermatogenic traits (DSP, DSPG) and 
spermatogenic apoptosis assessed by cell death detection ELISA in control, E. cob: 
endotoxin infused, or scrotal insulated (48 h) bulls are shown in Figure 3-8. 



48 



55 

50 



CD 


45 






CO 




c 


40 


o 




'.= 




E 


35 


Q- 




co 




2 


30 




25 




85 


, — . 




c 
F 


80 


c/> 


75 


3 




CL 


70 







ro 


65 


K 




en 


60 


« 




I 


55 




50 




45 




39 5 


U 




' — • 




cd 


39.0 






CO 








CD 

Q. 


38.5 


fr 









h- 


38.0 






o 




0) 




(T 


37.5 




4 



37.0 



• Control 
t Endotoxin 



Tx 



n; 



,h u. 





60 120 180 240 300 360 

Time (min) 



Figure 3-1 . Least squares means (± SEM) of the effect of E.coli endotoxin infusion (100 
ng/kg) on respiration rate, heart rate, and rectal temperature in bulls. Respiration rate and 
rectal temperatures were affected by time (P<0.05 respectively). 



90 - 



50 



Control 

Endotoxin 

SI 



49 




~~ i 1 1 1 1 <— 

3 6 8 



Time (days) 



Figure 3-2. Spermatozoal motility (Least squares means ± SEM) in control, E.coli 
endotoxin infused (100 ng/kg) (endotoxin), or scrotal insulated for 48 h (SI) bulls. 



50 



CD 

CD 



o 

c 
< 






CO 

CD 



ro 
E 

o 

c 

-Q 
< 

TO 
"D 

C 



CO 



90 - 



CD 


85 


o 




N 




O 


80 


to 




b 


75 


CI) 




Q 




CO 


/U 


CO 




F 


65 


i_ 




o 




z. 


60 



55 



6 - 




35 
30 
25 
20 
15 
10 
5 





-i 1 1 r 



Qbntrol 

Endotoxin 

SI 



t 1 1 r 





-i 1 1 1 — 

-3 



"i i i i 1 1 1 — 

3 6 8 



Time (days) 



Figure 3-3. Percentage of normal spermatozoa, primary abnormalities, and secondary 
abnormalities (Least squares means ± SEM) in control, E.coli endotoxin infused (100 
ng/kg) (endotoxin), or scrotal insulated for 48 h (SI) bulls. 



51 



Table 3-1. Effect of treatment (control, endotoxin, or scrotal insulation) on sperm 
motility, normal spermatozoa, primary abnormalities, and secondary abnormalities". 

Item Control Endotoxin Scrotal insulation 

Motility 

Day -3 62.5 ± 9.27 67.5 ±6.55 77.5 ±6.55 

Day 8 85.0 ±9.27 85.0 ±6.55 78.8 ± 6.55 

Difference (d -3 - d 8) -22.5 ± 9.86 -2.5 ± 6.97 8.8 ±6.97 

Normal spermatozoa b 

Day -3 74.5 ±10.06 2.0 ±8.21 79.9 ±7.11 

Day 8 81.0 ±10.06 73.7 ±8.21 70.3 ±7.11 

Primary abnormalities b 

Day -3 5.3 ±1.31 3.0 ±1.07 5.3 ± 0.93 

Day 8 6.5 ±1.31 5.0 ±1.07 4.5 ± 0.93 

Secondary abnormalities' 5 

Day -3 20.3 ± 9.49 25.0 ±7.75 14.9 ±6.71 

Day 8 12.5 ±9.49 21.3 ±7.75 25.3 ±6.71 

"Least squares means ± SEM. 

b Semen fixed with buffered isotonic formal-saline (FBS). 



52 



Table 3-2. Semen and testicular traits at 8 d in control, R. coli endotoxin or scrotal 
insulated (SI) treated bulls ab . 



Treatment 



Item 



Control 



Endotoxin 



SI 



Normal spermatozoa , % 
Primary abnormalities , % 
Secondary abnormalities , % 
Scrotal circumference, cm 
Testicular circumference, cm 
Paired testicular vol., cm 3 
Paired testicular wt, g 
Paired epididymal wt, g 



78.1 ±7.25 
15.5 ±6.30 
6.4 ±6.52 
37.0 ±0.88 
20.9 ±0.38 



72.4 ± 6.28 
14.2 ±5.46 
13.7 ±5.65 
35.9 ±0.62 
20.1 ±0.29 



78.2 ±6.28 

15.7 ±5.46 

5.8 ±5.65 

36.4 ± 0.62 

20.1 ±0.38 



1298.5 ±175.05 1 124.4 ± 135.60 1090.5 ± 175.05 

692.8 ±54.02 635.1 ±41.84 618.8 ±54.02 

71.0 ±5.49 75.8 ±4.25 63.2 ±5.49 



"Least squares means ± SEM. 

b There was no effect on semen and testicular traits. 

°Semen stained with negrosin and eosin. 






53 



Table 3-3. Sertoli cells, germ cells (spermatogonia A, preleptotene, pachytene, and round 
spermatid stage 8, and Sertoli:germ cells ratio in 20 seminiferous tubules stage VIII in 
control, R cob endotoxin infusion (100 ng/kg) (endotoxin), or scrotal insulated for 48 h 
(SI) bulls 3 . 



Item Control Endotoxin Scrotal insulation 



Sertoli cells 7.1 ± 1.80 8.4 ± 1.56 8.4 ± 1.56 

Germ cells 160.9 ± 13.20 169.0 ±11.43 175.6 ±11.43 
Sertoli: germ cell 22.7 ±3.78 23.0 ±3.27 22.5 ± 3.27 

Apoptotic cell 0.317 ±0.0969 0.413 ±0.0839 0.338 ±0.0839 



"Least squares means ± SEM. 



54 



=3 
.Q 

3 



CD 
O 



CD 

o 

o 
c 

CD 
CO 

CD 
CD 

TO 
i— 
<D 
> 
< 



-Q 



CD 
U 



CD 

o 



CD 

CD 



JD 

=1 

.a 



CD 
O 



TO 

a: 

"cd 
o 



CD 

a 

o 

tr 

CD 
CO 



10 - 






















8 - 












6 - 








4 - 
















2 - 
n 
















u 
180 - 




T 




160 - 
140 - 
120 - 
100 - 




T 


^m 
















80 - 
60 - 

40 - 












20 - 
n 
















u 
25 - 




T 




T 




T 




20 - 
















15 - 
















10 - 
















5 - 
- 








■..:■::■:.: km .::::: : :::- : 









Control Endotoxin SI 

Treatment 



Figure 3-4. Sertoli cells, germ cells (spermatogonia A, preleptotene, pachytene, and 
round spermatid 8), and Sertoli:germ cells ratio in 20 stage VIII seminiferous tubules 
(Least squares means ± SEM) in control, R co]i endotoxin infusion (100 ng/kg) 
(endotoxin), or scrotal insulated for 48 h (SI) bulls. 



55 









aT 






=3 


0.5 - 






_Q 








13 








-4— ' 










i2 


0.4 - 








"a3 










o 


0.3 - 


























_W3 




"53 














O 














o 


0.2 - 












'•4—* 














o 










: : 








+-- 


















O. 


















o 

Q. 


0.1 - 
















< 


0.0 - 




i 




I 




I 





Control Endotoxin SI 
treatment 



Figure 3-5. Apoptotic cells per stage VIII seminiferous tubule (Least squares means ± 
SEM) in control, E. coli endotoxin infusion (100 ng/kg) (endotoxin), or scrotal insulated 
for 48 h (SI) bulls. 



56 



13 

.o 

o 

I 

c 

E 
d) 



O 



o 

■*-> 
Q. 
O 
Q. 

< 



0.7 - 




T 




0.6 - 
0.5 - 
0.4 - 
0.3 - 






| 


- 




0.2 - 
0.1 - 
0.0 - 




_ 




ilililllll 




, 





Control Endotoxin 
Treatment 



SI 



Figure 3-6. Apoptotic cells per seminiferous tubule (LSMeans ± SEM) in control, E, coli 
endotoxin, and 48 h of scrotal insulation (SI). Spermatogenic apoptosis, assessed by 
Tunel staining, did not differ among treatments. 






57 



E 

c 

o 



Q 

o 

c 

CO 

■e 

o 
in 

< 



1.0 - 
0.8 



0.6 - 



0.4 - 
0.2 



0.0 - 




~l r 



~i 1 1 r 

10 20 30 40 50 60 70 80 



Concentration (ug) 






Figure 3-7. Effect of testicular concentration (n=3 bulls) on light absorbance at 405 nm, 
using a cell detection ELISA. There was an effect of concentration on light absorbance 
(P< 0.05), and orthogonal contrasts 2.5 vs 5, 10, 20, 40, and 80 /u,g, 5 vs 10, 20, 40, and 
80 Aig, and 10 vs 20, 40, and 80 /u.g were significant (P<0.05). 



58 



E 

g 2000 


i 1 S| 


)err™ 


atogenic 


Apoptosis 


-r- 




E 
















| 1500 

o 


- 














ro 1000 


- 














o 

c 

CD 

•e 500 

o 

w 
.O 

< 


- 














18 


- ■■" DSPG(xl0 6 ) 


p 16 


c=3 DSP ( xlO 9 ) T 


1 14 

2 12 

0_ 

E 10 


- 


|M 


8. 8 
% 6 

CD 4 
Q 

2 




1 


1 


L 



Control Endotoxin S.I. 

Treatment 



Figure 3-8. Least squares means (± SEM) on the effect of E.coli endotoxin infusion (100 
ng/kg) or 48 h of scrotal insulation (SI) on spermatogenic apoptosis, DSPG (daily sperm 
production/g) and DSP (daily sperm production). There was no effect of treatment on any 
of the parameters evaluated. 



59 

Spermatogenic traits, determined by DSP and DSPG, and spermatogenic apoptosis, were 
not different among groups. The in-situ 3' end-labeling of apoptotic cells with ApopTag 
showed staining of less than 1 cell per seminiferous tubule in either control or treated 
bulls, and no stained cells were observed in negative controls. 
Experiment 2 

Percent sperm motility (LSMeans ± SEM) on d -6, d -4, and d 1 (in relation to 
either d or initiation of treatment) is shown on Table 3-4 and Figure 3-9. Sperm 
motility did not differ as a result of treatment (control or 48 h of SI), breed, or time. Least 
squares means (± SEM) for FBS sperm morphology (percentage of normal spermatozoa, 
percentage of primary abnormalities, and percentage of secondary abnormalities) are 
shown on Table 3-4 and Figure 3-10 and Figure 3-11. The percentage of normal 
spermatozoa was not affected by treatment or time. However, percentage of normal 
spermatozoa was affected by breed (P<0.01). Angus bulls had less normal spermatozoa 
when compared with Romosinuano and Senepol (P<0.01). Primary abnormalities were 
not affected by treatment or breed. However, primary abnormalities were influenced by 
time (P<0.05). Primary abnormalities, regardless of treatment or breed, were higher at d 
1 (PO.01) when compared with primary abnormalities at d -6 or d -4. Secondary 
abnormalities were not influenced by treatment or time. However, there was an effect of 
breed on secondary abnormalities (PO.001). Secondary abnormalities were higher 
(PO.01) in Angus bulls than in Romosinuano and Senepol bulls. 



60 



Table 3-4. Sperm motility, normal spermatozoa, primary abnormalities, and secondary 
abnormalities in Angus, Romosinuano, and Senepol bulls 3 . 



Item 



Angus 



Romosinuano 



Senepol 



Motility 

Day -6 77.5 ± 7.04 77.4 ± 8.30 

Day -4 68.8 ± 7.04 79.0 ± 8.30 

Day 1 75.6 ± 7.04 79.9 ± 8.30 

Difference b -2.6 ± 4.43 -1.9 ± 5.42 

Normal spermatozoa 

Day -6 67.9 ±3.06 84.9 ±4.33 

Day -4 66.1 ±3.06 86.8 ±4.33 

Dayl 72.1 ±3.06 85.6 ±4.33 

Primary abnormalities' 1 

Day -6 10.1 ± 1.30 4.5 ±1.84 

Day -4 9.5 ±1.30 3.6 ±1.84 

Dayl 11.2 ±1.30 6.0 ±1.84 

Secondary abnormalities 

Day -6 21.9 ±2.90 10.6 ±4.10 

Day -4 24.4 ±2.90 9.6 ±4.10 

Dayl 16.7 ±2.90 8.4 ±4.10 

"Least squares means ± SEM.. 

b Difference on the average of d -6 and d -4 minus d 1 . 

°Orthogonal contrast, breed (Angus vs Romosinuano and Senepol; P< 0.05). 

d Orthogonal contrast, time (day 1 vs day -6 and day -4; P< 0.05). 



73.8 ±9.96 

68.8 ±9.96 
81.3 ±9.96 

-10.1 ±6.26 

86.3 ± 6.61 

84.3 ±6.61 
79.1 ±6.61 

4.4 ±1.54 
3.8 ±1.54 

10.4 ±1.54 

9.3 ± 3.42 

11.9 ±3.42 

10.5 ±3.42 



61 



85 

~ 80 

J 75 

o 

5 70- 

65 - 

60 



90 - 
2 80- 
j 70 - 

60 
50 



~i i 



T I 




—0— Control 

— O— Scrotal insulation 




# Angus 
— O— Romosinuano 
m Senepol 



n i i i r 



-4 -2 

Time (days) 



1 



Figure 3-9. Spermatozoal motility (Least squares means ± SEM) in control or scrotal 
insulated for 48 h (SI) Angus, Romosinuano, or Senepol bulls. 



62 




Time (days) 



Figure 3-10. Percentage of normal spermatozoa, primary abnormalities, and secondary 
abnormalities (Least squares means ± SEM) in control or scrotal insulated for 48 h (SI) 
bulls. 



63 




Time (days) 



Figure 3-11. Percentage of normal spermatozoa, primary abnormalities, and secondary 
abnormalities (Least squares means ± SEM) in Angus, Romosinuano, or Senepol bulls. 



64 



Table 3-5. Body weight, rectal, and scrotal temperatures in controls and treated bulls of 3 
breeds at 24 h following initiation of scrotal insulation (SI) in bulls 3 . 



Breed 



Item 



Angus 



Romosinuano 



Senepol 



No. of bulls 
Body weight (kg) 
Rectal Temp (°C) 

Control 

Scrotal insulated 

Control b (Log 10 ) 

Scrotal insulated b (Log 10 ) 
Scrotal Temp (°C) 

Control 

Scrotal insulated 

Control (Log 10 ) 

Scrotal insulated (Log 10 ) 



8 



419.3 ± 12.34" 534.8 ±15.11° 



39.2 ±0.26 
38.6 ±0.26 
1.6 ±0.003 
1.6 ±0.003 

31.2 ±0.55 
34.4 ±0.64 
1.49 ±0.01 
1.53 ±0.01 



38.1 ±0.36 
38.3 ±0.26 
1.6 ±0.005 
1.6 ±0.003 

30.8 ±0.78 

33.7 ±0.55 

1.5 ±0.01 

1.52 ±0.01 



"Least squares means ± SEM. 

"Orthogonal contrast, Angus vs Romosinuano and Senepol (P< 0.05). 

°Orthogonal contrast, Control vs scrotal insulated (P< 0.05). 



522.0 ±17.45° 

37.5 ±0.36 
38.0 ±0.36 
1.6 ±0.005 
1.6 ±0.005 

31.0 ±0.78 

32.7 ±0.78 

1.5 ±0.01 

1.5 ±0.01 



65 
Table 3-5 shows LSMeans (± SEM) for body weight (BW), rectal and scrotal 
temperatures 24 h after initiation of treatment. Body weight of Angus bulls was lower 
than Senepol and Romosinuano (P< 0.01), and rectal temperature was higher in Angus 
when compared to Senepol and Romosinuano bulls (P<0.01). However, rectal 
temperature was not affected by scrotal insulation. Scrotal temperature was not affected 
by breed, and scrotal temperature was higher in scrotal insulated than in control bulls 
(PO.05). 

Least squares means (± SEM) for the effect of treatment (control or 48 h of SI) 
and day of tissue harvest on semen and testicular traits are shown in Table 3-6. 
Percentage of secondary abnormalities were higher in both control and SI bulls at d 2 than 
at d 4 (P<0.05). In contrast, paired epididymal weight was lower in control than in SI 
bulls at d 2 and at d 4 (P<0.05). Percent of normal spermatozoa, percent of primary 
abnormalities, scrotal and testicular circumference, paired testicular volume, and paired 
testicular weight were not affected by treatment or day. Least squares means (± SEM) for 
the effect of breed (regardless of treatment) on semen and testicular traits are shown in 
Table 3-7. Semen characteristics and paired epididymal weight were not influenced by 
breed. Scrotal circumference, paired testicular volume, testicular circumference, and 
paired testicular weight were smaller in Angus bulls than in Senepol and Romosinuano 
(P<0.001 respectively). 

Results of the effect of scrotal insulation (SI) on Hsp70 levels (least squares 
means ± SEM) in bull semen are shown in Table 3-8 and Figures 3-12, 3-13, and 3-14. 
Heat shock protein 70 levels (/u-g/ml) were not influenced by breed or treatment (P>0.05). 



66 



Table 3-6. Semen and testicular traits in control or 48 h scrotal insulated (SI) bulls" 



Day 2 b Day 4 b 

Item Control SI Control SI 

Normal 

spermatozoa', % 80.9 ±7.75 73.2 ±6.93 72.4 ± 7.75 86.6 ±6.93 

Primary 

abnormalities , % 8.5 ± 5.73 10.5 ±5.13 21.6 ±5.73 8.0 ±5.13 

Secondary 

abnormalities cd , % 11.1 ±2.19 8.2 ±1.96 6.0±2.19 4.4 ± 1.96 

Scrotal 

circumference, cm 32.9 ±1.87 33.7 ±1.67 32.1 ± 1.87 34.1 ± 1.67 

Testicular 

circumference, cm 17.1 ±0.99 18.2 ±0.88 16.9 ±0.99 18.7 ±0.88 

Paired testicular vol., 

cm 3 723.0 ±101.16 701.2 ±90.48 544.7 ±101.16 826.4 ±90.48 

Paired testicular wt, 

g 426.8 ±74.56 486.4 ± 66.69 398.4 ±74.56 518.1 ±66.69 

Paired epididymal 

wt,g 48.4±7.84 b 62.4±7.02 c 42.5 ± 7.84 b 60.6 ±7.02° 



"Least squares means ± SEM. 
b In relation to day or at the beginning of treatment 
c Semen stained with nigrosin and eosin. 
d Contrast, time (day 2 vs day 4; P< 0.05). 



67 



Table 3-7. Semen and testicular traits in control and treated Angus, Romosinuano, and 
Senepol bulls at 24 h following initiation of scrotal insulation (SI) a . 



Breed 



Item 



Angus 



Romosinuano 



Senepol 



No. of bulls 8 

Normal spermatozoa b , % 72. 1 ± 5.40 

Primary abnormalities' 5 , % 17.5 ± 4.19 

Secondary abnormalitiesb b , 9.8 ± 1.53 
% 

Scrotal circumference , cm 30.0 ± 0.72 



Testicular circumference , 
cm 

Paired testicular vol. , cm 3 

Paired testicular wt°, g 

Paired epididymal wt, g 



16.2 ± 0.47 
533.7± 51.10 
334.9 ±31.22 

46.1 ± 4.81 



88.2 ±6.61 

7.5 ±5.13 
4.8 ±5.93 

35.9 ±0.88 

19.1 ± 0.58 
858.0 ±62.59 
573.7 ±38.23 

60.6 ± 5.89 



76.4 ± 7.63 
7.3 ±5.93 
6.2 ±2.16 

35.7 ±1.01 

18.8 ± 0.67 
783.1 ±72.27 
536.5 ±44.14 

55.5 ± 6.81 



"Least squares means ± SEM. 

b Semen stained with nigrosin and eosin. 

"Orthogonal contrast, breed (Angus vs Romosinuano and Senepol; P< 0.05). 



68 



Table 3-8. Heat shock protein 70 Og/ml) in semen of Angus, Romosinuano, and Senepol 
bulls". 



Item 


Day -6 


Day -4 


Day 1 


Breed 








Angus 


260.7 ± 68.48 


163.6 ±68.48 


137.9 ±75.46 


Romosinuano 


172.8 ±80.71 


173.0 ±80.71 


66.2 ± 80.71 


Senepol 


309.0 ±96.85 


181.5 ±96.85 


25.8 ±96.85 


Treatment 








Control 


288.1 ±68.00 


180.0 ±68.00 


43.4 ±73.14 


SI48h 


201.9 ±60.65 


159.9 ±60.65 


110.6 ±60.65 


Time b 


247.5 ± 47.82 


172.7 ±47.82 


76.7 ±49.00 



"Least squares means ± SEM. 

b Orthogonal contrast, time (day 1 vs day -6 and -4; P<0.05). 



69 



^ 300 

E 

X 200 
o 

N 

CL 
C/3 

^ 100 



-•— Control 
■O- SI 24 h 




-i 1 1 — 

-6 -4 



-i r 

-2 



1 



Time (days) 



Figure 3-12. Heat shock protein 70 (Hsp70, Aig/ml) in semen (least squares means ± 
SEM) over time in control or scrotal insulation ( SI 24 h). Days in relation to initiation of 
SI treatment. Heat shock protein 70 (Hsp70) in semen was influenced (P<0.05) by time 
but not by treatment. 



70 







Time (days) 



Figure 3-13. Heat shock protein 70 (Hsp70, //g/ml) in semen (least squares means ± 
SEM) over time (days in relation to initiation of SI treatment), regardless of treatment, in 
Angus, Romosinuano, and Senepol bulls. Heat shock protein 70 (Hsp70) in semen was 
influenced (P<0.05) by time but not by breed. 



71 



300 



1 250 





I 



1 r 



-4 -2 1 

Time (days) 



Figure 3-14. Heat shock protein 70 (Hsp70, Aig/ml) in semen (least squares means ± 
SEM) over time (days in relation to initiation of scrotal insulation treatment), regardless 
of breed and treatment. Heat shock protein 70 (Hsp70) in semen was influenced (P<0.05) 
by time only. 



72 



a 

-Q 

o 

I 

g 

E 


M 

15 
o 
o 

o 

Q- 
O 
Q. 
< 




d2 d4 

Time (days) 



Figure 3-15. Effect of 48 h of scrotal insulation (SI) on apoptotic cells per seminiferous 
tubule (LSMeans ± SEM) in tissue harvested immediately (d 2) following removal of 
scrotal insulation, or 2 days later (d 4). Spermatogenic apoptosis, assessed by Tunel 
staining, was not effected by treatment. However, there was a tendency for an effect of 
time (P= 0.06). 



73 
However, Hsp70 levels were affected by time (PO.05). Results of the number of 
apoptotic cells per siminiferous tubule after 48 h SI or control bulls are shown in Figure 
3-15. Spermatogenic apoptosis, assessed by Tunel staining, was not effected by 
treatment. 

Spermatogenic traits and spermatogenic apoptosis (LSMeans ± SEM) in control 
and 48 h SI bulls are presented in Figure 3-16. Scrotal insulation showed no effect on 
DSP, DSPG, and spermatogenic apoptosis. Least squares means (± SEM) for DSP, 
DSPG, and spermatogenic apoptosis in Angus, Romosinuano, and Senepol bulls are 
shown in Figure 3-17. Spermatogenic apoptosis and DSPG did not differ among breeds. 
However, DSP was lower (P<0.05) in Angus than in Senepol and Romosinuano bulls. 

Discussion 

In experiment 1 , rectal temperature, heart and respiration rates were affected by 
time of assessment, but not by treatment, indicating that E. coli endotoxin caused no 
observable clinical changes compared to control animals. These results suggested that 
100 ng/kg, of R cob endotoxin administered intravenously in the bull, were not sufficient 
to induce pyrexia, despite achieving such effects in the horse (MacKay and Lester, 1992; 
Hawkins et al., 1993). In this study, neither semen nor testicular traits were adversely 
affected by either treatment (R coli endotoxin or 48 h of scrotal insulation). These semen 
findings are not surprising as it has been previously shown that abnormal spermatozoa 
started to appear 12 d after 48 h of SI (Vogler et al., 1993), whereas in the present study 
semen was only studied up to 8 d after SI, as the main objective was to evaluate 



74 



I 

E 



E 
c 
lo 
o 

■* 



o 
c 
ra 

XI 

i_ 
o 
<n 
X) 

< 



CD 
O 



C 

o 
o 

■o 
2 

0- 



& 
1 



en 
o 



o 

=3 



ID 

Q. 
CO 

TO 

Q 



2000 



1500 



1000 



500 




d2 d4 

Time (days) 



Figure 3-16. Effect of 48 h of scrotal insulation (SI) on spermatogenic apoptosis 
(absorbance at 405 nm) (LSMeans ± SEM), daily sperm production/g (DSPG), and daily 
sperm production (DSP) in tissue harvested immediately (d 2) following removal of 
scrotal insulation, or 2 days later (d 4). There was no effect of treatment on any of the 
variables. 



75 



E. 

E 
c 

in 
o 



(D 

o 

c 

ra 

n 

t_ 
o 

(/) 

< 



X 

t 

o 

B 

=! 
T3 

2 
Q- 

E 

s. 

CO 

re 
Q 



o 



o 

■■o 

3 



CO 

s 



2000 



1500 



1000 



500 




Angus Romosinuano Senepol 
Breed 



Figure 3-17. Spermatogenic apoptosis (absorbance at 405 nm), daily sperm production/g 
(DSPG), and daily sperm production (DSP) in Angus, Romosinuano, and Senepol bulls 
(LSMeans ± SEM). There were no breed effects in spermatogenic apoptosis and DSPG, 
although DSP was influenced by breed (Angus vs Romosinuano and Senepol; P<0.05). 



76 
spermatogenic changes, which are generally detectable earlier. In this study, 
Sertolli:germ cell ratio was higher than the ratios reported (Berndtson et al., 1987; 
Berndtson and Igboeli, 1989). However, this study is reporting ratios based on crude 
counts, and ratios reported by Berndtson et al. (1987) and Berndtson and Igboeli (1989) 
are based on Abercrombie's corrected counts. Testicular characteristics in both 
experiments were considered to be within the ranges reported for B. taurus bulls in 
Florida with the exception of Angus bulls used in experiment 2, which had smaller body 
weights, scrotal and testicular measures than Angus bulls of similar age (Fields et al. 
1979; Chenoweth et al, 1996; Chase et al., 1997). 

Heat shock protein 70 has been reported to prevent protein denaturation as a result 
of heat insult. Results in the present study suggest that SI for 24 h did not significantly 
increase Hsp70 levels in semen, even when SI increased scrotal skin temperature to 
similar levels previously reported by Vogler et al. (1993). However, Hsp70 was lower at 
d 1 than at d -6 and d -4, suggesting that some stimulus other than treatment was 
responsible for decreasing Hsp70. Such a stimulus could be environmental temperatures 
or animal handling stress. It is possible that the presence of such a stressor increased the 
demand for Hsp70 resulting in its utilization and subsequent decreased of Hsp70 levels in 
semen, even when the synthesis of Hsp70 was also increased as result of the stressor. 
The result of this scenario could be a net decrease of Hsp 70 in ejaculated semen. 

Spermatogenic traits (DSPG and DSP) were not affected by any treatment in this 
study, and values were within previously reported ranges (Weisgold and Almquist, 1979; 
Amann, 1981; Wildeus and Entwistle, 1982; McCool, 1990; Chenoweth et al., 1994). 



77 
However, in experiment 2, DSP for Angus bulls was lower than for Senepol and 
Romosinuano, although DSPG was not different among breeds. This indicated that 
spermatogenic efficiency, or production of spermatozoa per unit (g) of testicular 
parenchyma, was not different among breeds, and that the difference in DSP was most 
probably due to smaller testicles in Angus bulls compared to Senepol and Romosinuano 
bull. The lack of effect encountered with quantitative spermatogenic traits was also not 
unexpected, because it has been reported that elevated temperature treatment (10 or 20 h 
of SI) decreased the number of type B spermatogonia 14 days after treatment (Ross and 
Entwistle, 1979). Spermatogenesis in the present study was determined by counting 
elongated spermatids (DSPG and DSP), and in order to observe a decrease in the number 
of elongated spermatids as a result of damage to spermatogonia by 48 h of SI, a longer 
period (more than the d 2, d 4 or d 10 used in the present study) would be necessary. 
Spermatogenic apoptosis, as assessed by histology, tunel stain, and cell death 
detection ELISA, was not influenced by either E. coh endotoxin infusion or 48 h of 
scrotal insulation in this study. Furthermore, overall levels detected with the ELISA test 
were lower than those previously reported for cattle, using the same method (Hingst and 
Blottner, 1995). Possible reasons for these relatively low levels of detectable apoptosis 
include the following. Firstly, there is the possibility that none of the treatments caused 
increased testicular apoptosis. This is regarded as unlikely as 48 h scrotal insulation, at 
least, has consistently produced levels of spermatogenic dysfunction compatible with 
degeneration and associated apoptosis (Ross and Entwistle, 1979; Vogler et al., 1993). 
In the present study, scrotal temperatures were significantly higher, after 24 h of scrotal 



78 
insulation treatment, in treated than in control bulls. Also, the skin scrotal temperatures 
achieved in this study are within the ranges reported in the literature (Austin et al., 1961; 
Ross and Entwistle 1979; Entwistle, 1983a; Sidibe et al., 1993; Vogler et al., 1993). 
Although relatively few cells stained positive for apoptosis using the ApopTag 
peroxidase, this was in agreement with low apoptotic levels detected by the cell death 
detection ELISA. Secondly, it is possible that apoptosis assessment was done before, or 
after, apoptotic cells had been phagocytosed and rendered undetectable (Richburg and 
Boekelheide, 1996; Lin et al., 1997). However, in a more recent study, when a GnRH 
antagonist was used in the rat, testicular apoptosis began to increase after 5 days of 
treatment, reaching its peak 14 days after treatment (Hikim et al., 1997), suggesting that 
spermatogenic apoptosis assessment after 48 h of SI, in the bull, should be done 
following 8 days and before 14 days of treatment. This is because, in the bull, the 
spermatogonial population has been shown to have declined by d 14 after SI (Ross and 
Entwistle, 1979). 

This series of experiments failed to detect increased testicular apoptosis in bulls 
after E. coh endotoxin infusion or 48 h of scrotal insulation at 10 d after initiation of 
treatment or on d 2 or d 4 after initiation of 48 h of SI, suggesting either that these 
treatments caused no detectable changes in spermatogenic apoptosis, that the detection 
windows used were inappropriate, or that damage to cells as result of SI or E. coli 
endotoxin undergo a different type of cell death from apoptosis (Young et al., 1997). 
Also, the levels of spermatogenic apoptosis observed corresponded to basal levels of 
apoptosis, resulting probably from spontaneous degeneration of germ cells, that undergo 



79 
apoptosis. Since in the rat, it has been shown that germ cells that die during normal 
spermatogenesis die through apoptosis, suggesting that a basal level of spermatogenic 
apoptosis is always present during normal spermatogenesis (Bianco-Rodriguez and 
Martinez-Garcia, 1996). 



CHAPTER 4 

EFFECT OF DIETS CONTAINING FREE GOSSYPOL AND VITAMIN E ON 

SPERMATOGENESIS AND SPERMATOGENIC APOPTOSIS IN YOUNG 

HOLSTEIN BULLS 

Introduction 



Exposure of bulls to elevated temperatures whether environmental or 
experimental, results in decreased semen quality (Johnston et al., 1963; Fields et al., 
1979; Meyerhoeffer et al., 1985; Chase et al., 1993) and reduced spermatogenesis 
(Skinner and Louw, 1966). The deleterious effect of elevated temperature on 
spermatogenesis could be as a result of increased levels of reactive oxygen species in 
testicular tissue. Reactive oxygen species (ROS), or free radicals, have been proposed as 
major contributors to cell damage resulting from heat shock by causing lipid 
peroxidation, protein denaturation, impairment of the cytoskeleton, and disruption of 
calcium metabolism (Loven, 1988). Gossypol, a toxic polyphenols pigment produced in 
the pigment glands of roots, leaves, stems, and seeds of the cotton plant (Berardi and 
Goldblatt, 1969) has been associated with reproductive problems in several species 
including humans, rodents, and cattle (Liu, 1957; Liu and Segal, 1985, Jimenez et al., 
1989; Randel et al., 1992; Chase et al., 1994; Chenoweth et al., 1994), possibly by 
increasing free radical production (de Peyser et al., 1984; Barhoumi and Burghardt, 
1996). Since gossypol has been reported to increase reactive oxygen species and reduce 

80 



81 
antioxidants in hepatocytes and testes in the rat (Bender et al., 1988; Barhoumi and 
Burghardt, 1996), it is logical to suggest that antioxidants could counter the oxidative 
damage of gossypol on spermatogenic tissue in bulls. This concept was supported in a 
recent study where the spermatoxic effects of gossypol in the bull were countered by 
administration of the antioxidant, vitamin E (Velasquez-Pereira et al., 1995). 

In ruminants, feeding diets containing gossypol resulted in decreased 
spermatogenesis, possibly as a result of damage to germ cells within the germinal 
epithelium (Randel et al., 1992). In turn, these gossypol damaged germ cells could be 
removed via apoptosis, resulting in increased levels of spermatogenic apoptosis as a 
result of gossypol toxicity. 

The objective of this study was to evaluate the effect of long term feeding 
gossypol in cottonseed meal on testicular and spermatogenic traits as well as on 
spermatogenic apoptosis levels in young Holstein bulls. An additional objective was to 
determine whether the antioxidant, vitamin E, could counteract the spermatoxic effect of 
gossypol on testicular and spermatogenic traits and spermatogenic apoptosis which might 
otherwise be associated with gossypol induced cellular damage. 

Materials and Methods 

Materials 

Reagents were obtained as follows: Merthiolate, butanol, hydrogen peroxide 
(30%), and methyl green from Sigma Chemical Co. (St. Louis, MO); sodium chloride, 
sodium acetate, Permount, xylene, and Triton X-100 from Fisher Scientific (Fair Lawn, 



82 
NJ); Dulbeco's phosphate-buffered saline (DPBS) from Life Technologies Inc. (Grand 
Island, NY); cell death detection ELISA-kits from Boehringer Mannheim Co. 
(Indianapolis, IN); in-situ Tunel staining kits (ApopTag peroxidase S7101) from Oncor 
(Gaithesburg, MD); ethanol from AAPER Alcohol and Chemical Co. (Shelbyville, KY), 
and acetic acid from Scientific Products (McGaw Park, IL). 
Experimental Design 

Young Holstein bulls (n=24; 6 mo of age) were assigned to the following three 
isocaloric and isonitrogenous dietary groups (each n=8) that satisfied animal requirements 
for all other nutrients (NRC, 1989); 1) control (CONT), received a supplement containing 
soybean meal (SBM), corn and 30 IU of vitamin E/kg, 2) gossypol (GOSS), received a 
supplement containing CSM, corn and 30 IU vitamin E/kg, and 3) gossypol and vitamin 
E (G+VITE), received a supplement containing CSM, corn and 4,000 IU vitamin 
E/bull/d. Supplements GOSS and G+VITE were formulated to supply 14 mg of free 
gossypol/kg body weight (BW)/d (Table 4-1 and 4-2). Animals were housed in 12 pens, 
2 animals/pen and 4 pens/treatment, from 6 to 15 months of age. Animals had access to 
low quality hay in which the vitamin E concentration was less than 9 IU/kg. Supplements 
were recalculated monthly to ensure that the amount of free gossypol provided on an 
individual basis was 14 mg/kg BW/d. 

Semen was collected 2 weeks prior to sacrifice by electro-ejaculation, using a 
Lane IIIZ Pulsator (Lane Manufacturing, Denver, CO) in conjunction with a 3 ventral- 
electrode rectal probe. Immediately after semen collection, a smear stained with 



Table 4-1. Initial and final composition of dietary supplements 3 



83 







CONT 




GOSS 


G+VITE 


Item 


Initial 


Final 


Initial 


Final 


Initial 


Final 


Offered (kg/d) b 


2.6 


5.6 


2.7 


6 


2.7 


6 


DM (%) 


88 


88 


88 


88 


88 


88 


Ingredient b 














SBM (%) 


59 


71 


— 


— 


— 


— 


CSM (%) 


— 


— 


67 


80 


67 


80 


Corn (%) 


38 


27.5 


30 


18.5 


30 


18.5 


Limestone (%) 


1 


0.5 


1 


0.5 


1 


0.5 


Minerals (%) 


2 


1 


2 


1 


2 


1 


Vit. E (IU/kg) 


30 


30 


30 


30 


1481.5 


666.6 


Analyses 














(+)-gossypol (%) c 








0.33 


0.4 


0.32 


0.4 


(-)-gossypol (%) c 








0.79 


0.98 


0.77 


0.97 


Free gossypol (%) d 








0.08 


0.11 


0.08 


0.11 


Total gossypol (%) d 








1.06 


1.16 


1.06 


1.14 



a CONT = soybean meal (SBM) + corn + 30 IU E/kg; GOSS = cottonseed meal (CSM) + 

corn + 30 IU vitamin E/kg; G+VITE = CSM + corn + 4,000 IU vitamin E/animal/d. 

b As fed basis. 

c As fed. HPLC procedure (Calhoun et al., 1995; Kim and Calhoun, 1995) done at Texas 

A&M. 

d Asfed. AOACS procedure. Texas A&M. 



84 



Table 4-2. Average composition of dietary supplements 3 



Item 



Supplement 



CONT 



GOSS 



G+VITE 



Average values 
(+)-gossypol (%) 
(-)-gossypol (%) 
Free gossypol (%) 
Total gossypol (%) 
Vitamin A (IU/kg) d 
Vitamin E (IU/kg) b 
CP(%) e 
Ca (%) e 
K (%) e 
Mg(%) e 
P (%) e 
Cu (mg/kg) e 
Zn (mg/kg) e 
Mn (mg/kg) e 
Fe (mg/kg) e 
Se (mg/kg) e 



0.0 ± 0.00 
0.0 ±0.00 
0.0 ±0.00 
0.0 ±0.00 
2076.6 
38.8 ± 11.55 
38.6±4.31 
0.5 ±0.17 
1.4 ±0.44 
0.2 ± 0.06 
0.7 ±0.08 
16.7 ±7.65 
69.3 ±28.39 
52.6 ±26.95 
182.6 ±91.51 
0.2 ±0.11 



0.3 ± 0.03 
0.8 ±0.08 
0.1 ±0.01 
1.0 ±0.07 

2759.1 
44.7± 13.11 
37.9 ±2.03 
0.5 ±0.13 
1.4±0.10 
0.6 ±0.06 
1.1 ±0.05 
16.0 ±2.60 
73.3 ±11.77 
36.2 ±13.51 
161.7 ±76.54 
0.2 ± 0.04 



0.3 ±0.03 
0.8 ±0.08 
0.1 ±0.01 

1.0 ±0.07 
2759.1 

741.4 ±182.90 
37.1 ±2.43 

0.4 ±0.12 
1.42 ±0.36 

0.6 ±0.10 

1.1 ±0.08 
14.9±3.17 
70.5 ±13.85 
31.1 ±13.55 

139.0 ±40.00 
0.2 ± 0.04 



a CONT = soybean meal (SBM) + corn + 30 IU E/kg; GOSS = cottonseed meal (CSM) + 

corn + 30 IU vitamin E/kg; G+VITE = CSM + corn + 4,000 IU vitamin E/animal/d. 

b As fed basis. 

°Mean of 8 mixing ± SD. 

d As fed. A composited sample from all mixing dates. An IU/kg = Retinol acetate (ug/kg) 

* 2.91 

e DM basis. 



85 
nigrosin-eosin (Society for Theriogenology, Hastings, NE) was prepared. A minimum of 
200 sperm were assessed for morphology using oil-immersion bright-field microscopy (x 
1000) (Chenoweth et al., 1994). Abnormal sperm were classified based on the region 
(head, acrosome, midpiece, or tail) where the lesion was present (Chenoweth et al., 1994). 
Proximal droplets, abnormal acrosomes (e.g., knob), coiled tails, abnormal heads, and 
abnormal midpieces were categorized as primary sperm abnormalities, and distal 
cytoplasmatic droplets, kinked tails, and detached heads were categorized as secondary 
sperm abnormalities. 

Animals were sacrificed at 1 5 months of age using standard industry procedures at 
the University of Florida Meats Laboratory (Gainesville, FL). Testes were collected and 
immediately transported on ice to the laboratory. Each spermatic cord was trimmed, and 
the intact testis and tunic were weighed. The tunic was then removed, and both the tunic 
and testis were weighed separately. Testicular circumference was measured, and testis 
length, depth, and width were obtained to determine testicular volume. Paired testicular 
volume was calculated as the sum of the volume of the right and the left testes. Testes 
were considered as paraboloids, and volume was calculated using the equation v = irr^h, 
where r = (width + depth) / 4 and h = length (Chase, et al., 1997). Epididymides were 
removed, weighed, and mid-parenchymal sections were obtained for in-situ "Tunel" 
staining, spermatogenesis quantification (5 g), and for ELISA (1 g). 

The in-situ "Tunel" staining method used was the ApopTag S7101 kit (Hikim, et 
al., 1995). This method is based on the specific binding of terminal deoxynucleotidil 
transferase (TdT) to the newly formed 3'-OH of cleaved DNA. Testicular tissues were 



86 
fixed in 10% neutral buffered formalin for 24-36 h and paraffin-embedded and sectioned. 
Sections were de-paraffinized by two washes with xylene, two washes with absolute 
ethanol, one wash with 90% ethanol, one with 70% ethanol, and one with DPBS. 
Endogenous peroxidase was quenched using 2% hydrogen peroxide in DPBS, followed 
by two rinses with DPBS. Slides were placed in equilibrium buffer and then in working 
strength TdT enzyme. The reaction was stopped by adding working-strength stop/wash 
buffer. Two drops of anti-digoxigenin-peroxidase were applied to slides, and the 
peroxidase was detected with diaminobenzidine. Negative controls were established by 
adding distilled water instead of TdT enzyme during the preparation of working-strength 
TdT. Sections were counter-stained with 0.5% methyl green (w:v in 0.1 M sodium 
acetate, pH 4) and mounted with Permount. Cells stained positive for apoptosis were 
counted in ten seminiferous tubules epithelium/slide/bull. 

Aliquots of homogenized testicular parenchyma in DPBS were used for 
quantification of spermatogenic apoptosis using a cell death detection ELISA-kit (Hingst 
and Blottner, 1995). The cell death detection ELISA is a quantitative sandwich-enzyme- 
immunoassay that uses monoclonal mouse antibodies against DNA and histones that 
permit determination of mono- and oligonucleosomes in the cytoplasmic fraction of cell 
lysetes. Testicular parenchyma (1 g in 2 ml DPBS) was minced, freeze/thawed 3 times, 
homogenized for 1 min using a Polytron (Brinkmann Instruments, Westbury, NY), 
sonicated for 1 min at 4° C and then centrifuged at 25000 g for 30 min at 4° C. The 
supernatant was collected and stored in 100 ul aliquots at -20° C. The microtiterplate- 
modules (MM) were coated with 100 ul of coating solution containing the anti-histone 



87 
antibody, and cultured overnight at 4° C. Incubation buffer 200 ul were pipetted into 
each well, covered with plate cover foils, and incubated at room temperature (18° to 25° 
C) for 30 min to saturate the non-specific binding sites of the MM. After incubation, the 
incubation buffer was removed, and MM were washed three times with 300 ul washing 
solution. Then 100 ul of each sample solution containing 10 ug of testis equivalent/ 100 
ul were added to the MM, covered, and incubated for 90 min at room temperature. For 
background determination, 100 ul of incubation buffer were pipetted into three wells. 
Then, 100 ul of anti-DNA-peroxidase were added to the MM and incubated for 90 min at 
room temperature. After incubation, ABTS® (Boehringer) substrate was added to the 
wells, incubated at room temperature for 1 min, and the absorbance read at 405 nm. 

Quantitative spermatogenesis was assessed by counting elongated spermatids to 
determine daily sperm production (DSP) and DSP/g (DSPG) of testicular parenchyma 
(Chenoweth et al., 1994). Briefly, 5 g of testicular parenchyma were thawed, finely 
minced, and homogenized for approximately 1 min in 25 ml of working solution (0.9% 
NaCl, 0.05% Triton X-100, and 100 ppm Merthiolate diluted 1 :4 with distilled water). 
Then, 175 ml of working solution were added, mixed for 1 min, allowed to settle for at 
least 1 h, and then thoroughly mixed using a stirrer. Numbers of spermatozoa and 
elongated spermatids were counted using a hemocytometer, and the values were used to 
determine DSPG and DSP using the formula: 

DSPG = AX (B+Y) / (Time divisor)Y 

DSP = DSPG (0.99Z) 

Where X = hemocytometer count, Y = parenchyma sample weight, Z = testis parenchyma 



88 
weight, A = hemocytometer constant, B = dilution factor, and time divisor (5.32 = time 
divisor for Bos taurus) (Amann et el., 1974). 
Statistical Analysis 

Data were analyzed by least squares ANOVA, and means were separated by 
Duncan multiple range test, both using the PROC GLM procedure of SAS (1989). 
Testicular and semen characteristics as well as spermatogenic apoptosis were analyzed as 
a completely randomized design. Treatment and pen within treatment effects were tested 
using bull within pen and treatment as an error term. The pen within treatment effect was 
removed from the final model because it was not significant (Pk 0.2). All data are 
presented as least squares means ± SEM (LSMeans ± SEM). 

Results 

Least squares means (± SEM) for semen and testicular characteristics are shown 
in Table 4-3. The percentage of normal spermatozoa and the percentage of spermatozoa 
with primary sperm abnormalities were affected by treatment (30±7.0vs68±6.7 and 55 
± 6.0 %, and 59 ± 6.0 vs 24 ± 6.0 and 38 ± 5.2 %, for GOSS, CONT, and G+VITE 
respectively; PO.05). In contrast, no treatment effect was observed on secondary sperm 
abnormalities, paired testicular and epididymal weight, or scrotal circumference. 

Spermatogenic apoptosis, assessed by Tunel staining (LSMeans ± SEM), is shown 
in Figure 4-2. The number of apoptotic cells per seminiferous tubule did not differ as 
result of treatment (0.43 ± 0.05, 0.51 ± 0.05, and 0.52 ± 0.05 for CONT, GOSS, and 
G+VITE respectively). 



Table 4-3. Sperm morphology and testicular traits in young Holstein bulls fed diets 
containing gossypol, gossypol plus vitamin E, and controls 3 



89 







Supplement b 




Item 


CONT 


GOSS 


G±VITE 


Normal, % 


68 ± 6.7 d 


30 ±7.0° 


55 ± 6.0 d 


Primary abnormalities , % 


24 ± 6.0 d 


59 ±6.0° 


38±5.2 d 


Secondary abnormalities , % 


8 ±2.0 


11 ±2.0 


6± 1.8 


Scrotal circumference, cm 


32 ±0.8 


31 ±0.8 


31± 0.8 


Paired testicular wt, g 


495 ± 37.0 


490 ± 37.0 


564 ±37.0 


Paired testicular vol., cm 3 


707 ±45.0 


657 ±45.0 


748 ± 45.0 


Paired epididymal wt, g 


48±3.1 


42 ±3.1 


51±3.1 



"Least squares means ± SEM. 

b CONT = soybean meal (SBM) + corn + 30 IU E/kg; GOSS = cottonseed meal (CSM) + 

corn + 30 IU vitamin E/kg; G±VITE = CSM + corn + 4,000 IU vitamin E/animal/d. 

Trimary = proximal droplet, abnormal acrosome, coiled tail, abnormal head, and 

abnormal midpiece. Secondary = distal cytoplasmic droplet, kinked tail, and detached 

head. 

de Within rows, LSMeans ± SEM with different superscript are significantly different (P< 

0.05). 



90 



JO 

-O 

-t-< 
U5 

2 

i 

c 

'E 

Q) 

O 
o 

o 

a. 
o 

Q. 
< 



0.6 - 












T 




0.5 - 
0.4 - 


H ■ 






0.3 - 












0.2 - 








■n 








0.1 - 
















0.0 - 




I 




i 




i 





Control Goss G+VitE 

Treatment (Diet) 



Figure 4-1 . Apoptotic cells per seminiferous tubule in bulls fed gossypol (GOSS), 
gossypol plus vitamin E (G+VITE), and controls (CONT). Spermatogenic apoptosis, 
assessed by Tunel staining, did not differ between groups. 



91 
Sperm production, described as DSPG and DSP (LSMeans ± SEM), is shown in 
Figure 4-2. Daily sperm production/g (10.2 ± 1.0 vs 14.6 ± 1.0 and 17.6 ± 1.0 xlO 6 
spermatozoa for GOSS, CONT, and G+VITE respectively; P<0.05) and DSP (2.2 ± 0.3 
vs 3.2 ± 0.3 and 4.1 ± 0.3 xlO 9 spermatozoa for GOSS, CONT, and G+VITE 
respectively; P<0.05) were lower in gossypol alone (GOSS) fed bulls compared with 
either control bulls (CONT) or those fed both gossypol and 4000 IU of vitamin E 
(G+VITE). Least squares means ± SEM for spermatogenic apoptosis in the three groups 
are also shown in Figure 4-2. Gossypol had no effect on spermatogenic apoptosis (696.9 
± 13.21, 721 ± 14.98, and 703.3 ± 14.01 mU/mg for CONT, GOSS, and G+VITE 
respectively). However, there was an inverse relationship between sperm production 
(both DSPG and DSP) and the level of spermatogenic apoptosis (r=-0.46; PO.05 and r=- 
0.55; PO.05, DSPG and DSP respectively). 

Discussion 

Bulls fed diets containing gossypol alone showed a lower percentage of normal 
spermatozoa and higher percentage of primary sperm abnormalities compared to bulls in 
CONT and G+VITE groups. Similar results were reported by Chenoweth et al. (1994) 
where bulls fed similar levels of free gossypol showed a significant reduction in the 
number of normal spermatozoa and a higher percentage of midpiece abnormalities than 
did controls. However, in other reports, primary sperm abnormalities were not affected 
when bulls were fed diets containing gossypol (Jimenez et al., 1989; Chase et al., 1994). 



92 



CD 

E 
D 
E 

E 

c 

m 
o 

-t— ' 

03 

(I) 
O 

c 

03 

-9 
o 

I 



c 
o 

o 

13 
T3 
O 



CO 



800 
700 
600 
500 
400 
300 
200 
100 



20 



15 



10 



Apoptosis 



DSPG (x10 D ) 
DSP (x10 9 ) 




Control Goss G+VitE 

Treatment (Diet) 



Figure 4-2. Levels of spermatogenic apoptosis and sperm production in bulls fed 
gossypol (GOSS), gossypol plus vitamin E (G+VITE), and controls (CONT). 
Spermatogenic apoptosis, assessed by cell death detection ELISA did not differ between 
groups, but sperm production, assessed as daily sperm production per gram of 
parenchyma (DSPG) and daily sperm production (DSP), were lower (P<0.05) in GOSS 
than control or G+VITE. 



93 

In contrast, in the present study, bulls fed control (CONT) diets and diets containing both 
gossypol and vitamin E (G+VITE) showed significantly higher levels of normal 
spermatozoa and lower levels of primary sperm abnormalities than bulls fed gossypol 
alone (GOSS). This suggests that dietary vitamin E was able to protect the spermatogenic 
epithelium of bulls from the spermatoxic effect of gossypol. 

In the present study, gossypol did not adversely affect testicular traits (paired 
testicular and epididymal weights, paired testicular volume, and scrotal circumference). 
However, bulls supplemented with both gossypol and vitamin E showed a tendency to 
have larger testicular traits (paired testicular and epididymal weights, paired testicular 
volume, and scrotal circumference) than either control bulls or those fed with gossypol 
alone. In contrast, bulls supplemented with gossypol alone showed a tendency to have 
lower testicular traits. A similar tendency of lower, albeit non significant, paired 
testicular weights have been previously reported for bulls fed gossypol when compared 
with controls (Chase et al., 1994; Chenoweth et al., 1994), and results in this study are in 
agreement with previous findings showing that scrotal circumference and testicular 
weights were not adversely affected by gossypol (Jimenez et al., 1989; Chase et al., 
1994). 

Daily sperm production (DSP) and daily sperm production per g of testicular 
parenchyma (DSPG) were lowered in bulls supplemented with gossypol alone (GOSS) 
than in control (CONT) bulls and those supplemented with gossypol plus vitamin E 
(G+VITE). Daily sperm production and DSPG in CONT and G+VITE supplemented 
bulls were within values previously reported for dairy bulls (Amann et al., 1976; Amann, 



94 
1981). Lower DSPG and DSP have been previously reported in bulls fed gossypol 
(Chenoweth et al., 1994). This lower sperm production in GOSS bulls probably occurred 
via damage to the spermatogenic epithelium, as previously suggested (Randel et al., 
1992). Also, since gossypol has been reported to cause damage to the spermatogenic 
epithelium by reducing the number of germinal cell layers without reducing tubule 
diameter (Chase et al., 1994), the lack of change in both scrotal circumference and 
testicular weights in gossypol treated bulls in this study might be expected. 

Surprisingly, gossypol treatment alone had no effect on estimated levels of 
spermatogenic apoptosis (Tunel staining or ELISA), despite significantly reducing sperm 
production. However, there was a negative correlation between sperm production (DSPG 
and DSP) and spermatogenic apoptosis. Such a relationship concurs with results obtained 
during testicular involution in seasonal breeders (Blottner et al., 1995; Hingst and 
Blottner, 1995), supporting the underlying hypothesis that increased spermatogenic 
apoptosis is associated with decreased sperm production. Failure to detect differences in 
spermatogenic apoptosis in this experiment could result from the following 
considerations; a) the number of bulls studied was too small, b) spermatogenic cells 
damaged by gossypol, as reflected in decreased spermatogenesis, might undergo a form of 
cell death different from apoptosis (Young et al., 1997), or c) methods used in this study 
were inadequate to detect a difference in levels of spermatogenic apoptosis among 
groups. 

In defense of the current procedures, the number of bulls used in this experiment 
was sufficient to detect significant differences in DSPG and DSP among groups. In 



95 
addition, spermatogenic apoptosis has been reported to increase in other species after 
exposure to hormonal imbalance, spermatoxicity, or increased testicular temperature 
(Shikone et al., 1994; Richburg and Boekelheide, 1996; Hikim et al., 1997). Also, 
methods used to assess spermatogenic apoptosis in this study have been previously 
successfully employed in testicular tissue. The ELISA kit used to quantify spermatogenic 
apoptosis was similar to one previously used to determine spermatogenic apoptosis in 
bulls (Hingst and Blottner, 1995). Despite this, spermatogenic apoptosis levels detected 
in this study were lower than those reported by Hingst and Blottner (1995). The in-situ 
"Tunel" kit (ApopTag S7101) was similar to one previously used in testicular tissue in 
the rat (Hikim et al., 1995). Also, in-situ "Tunel" stained slides from our laboratory were 
validated by Oncor technicians, providing assurance that the methodology and reagents 
were satisfactory. 

In conclusion, free dietary gossypol had an adverse effect on percentage of normal 
sperm, percentage of primary abnormalities, DSPG, and DSP in young Holstein bulls, 
although testicular traits and spermatogenic apoptosis were not affected. Vitamin E, fed 
at 4,000 IU vitamin E/animal/d, was able to counteract the spermatoxic effects of 
gossypol, suggesting that vitamin E supplementation in the diets of bulls fed cottonseed 
meal could allow this valuable feedstuffs to be fed safely to puberal bulls. 



CHAPTER 5 
BREED EFFECTS ON SPERMATOGENESIS AND SPERMATOGENIC APOPTOSIS 

IN BULLS 

Introduction 



High environmental temperatures and high humidity, as encountered in 
subtropical regions of the United States, have been associated with lower fertility of cattle 
(Dunlap and Vincent, 1971; Badinga et al., 1985 ). Although high environmental 
temperatures can cause an increase in early embryonic mortality (Ealy et al., 1993), the 
bull probably contributes to this problem (Igboeli and Rakha, 1971; Wildeus and 
Entwistle, 1983), via decreased semen quality during the summer months (Johnston et al., 
1963; Igboeli and Rakha, 1971; Fields et al., 1979; Wildeus and Entwistle, 1983; Chase 
et al., 1993). Lowered semen quality has also been reported to occur after experimental 
exposure of bulls to elevated temperatures (Johnston et al., 1963; Meyerhoeffer et al., 
1985). Breed of sire may also influence both pregnancy rates and calf survival in 
subtropical environments (Gonzales-Padilla et al., 1969; Crockett et al., 1973). Also, 
breed differences in semen quality after exposure to elevated temperatures have been 
reported (Igboeli and Rakha, 1971; Rhynes and Ewing, 1973; Fields et al., 1979; Wildeus 
and Entwistle, 1983; Chase et al., 1993; Wildeus and Hammond, 1993). Differences in 
spermatogenic response to elevated temperatures between Bos taurus and Bos indicus 



96 






97 
genotypes most likely reflect wider aspects of environmental adaptability which 
differentiate between these groups (Kumi-Diaka et al., 1981 ; Kamwanja et al., 1993). 

Tropically adapted Bos taurus breeds such as Senepol (N'Dama x Red Poll 
crossbreed originated in St. Croix, U.S.V.I.) and Romosinuano (criollo-type native of 
Colombia) have been reported to be heat tolerant (Hammond et al., 1995), as well as Tuli, 
a Sanga breed (tropically adapted) that originated in Africa. Also, crossbreds such as the 
Senepol x Angus (SA) and Tuli x Angus (TA) have been reported to be heat tolerant 
similarly to the Brahman x Angus (BA) (Hammond et al., 1998). Rectal temperatures, 
during the Florida summer, have been observed to be lower in Senepol and Romosinuano 
than in Angus bulls (Chase et al., 1993; Hammond 1996). Also, semen quality has been 
reported to be lower during the summer in Angus and in Holstein bulls when compared 
with Senepol bulls (Chase et al., 1993; Wildeus and Hammond, 1993). This suggests that 
spermatogenesis in tropically adapted breeds such as Senepol, Romosinuano, and 
Brahman could be more efficient than in temperate breeds such as Angus, during periods 
of elevated ambient temperatures such as during the summer in Florida. 

Depression of spermatogenesis following exposure to elevated temperatures has 
been reported in the bull (Skinner and Louw, 1966; Ross and Entwistle, 1979; 
Meyerhoeffer et al., 1985; Vogler et al., 1993). Such spermatogenic depression is quite 
possibly associated with increased spermatogenic apoptosis (or programmed cell death). 
Evidence to support the idea that spermatogenic apoptosis in the bull could be increased 
by exposure to elevated temperatures can be found in the rat. In the adult rat testis, 
spermatogenic apoptosis was reported to increase following elevation of testicular 



98 
temperature due to cryptorchidism (Shikone et al., 1994). Spermatogenic apoptosis has 
been reported to occur in bulls, in seasonal breeders, in hamsters, in rats, in humans, and 
others (Shikone et al., 1994; Hingst and Blottner, 1995; Hikim et al., 1997; Lin et al., 
1997). 

The objectives of this study were 1) to evaluate the effect of breed type on 
testicular traits and extragonadal sperm reserves (ESR) among tropically adapted B. 
taurus and B. indicus (Romosinuano, Brahman, and Nelore x Brahman) and temperate B, 
taurus (Angus) bulls; 2) to determine the effect of breed type on testicular and 
spermatogenic traits and spermatogenic apoptosis in young tropically adapted crossbred 
bulls during the summer in central Florida (SA, TA, and BA); and 3) to obtain more 
information on testicular and spermatogenic traits and spermatogenic apoptosis in 
tropically adapted Miniature Brahman and Romosinuano bulls during the summer in 
central Florida. 

Materials and Methods 

Materials 

Merthiolate was purchased from Sigma Chemical Co. (St. Louis, MO). Sodium 
chloride, and triton X-100 were purchased from Fisher Scientific (Fair Lawn, NJ). 
Dulbeco's phosphate-buffered saline (DPBS) was purchased from Life Technologies Inc. 
(Grand Island, NY), and cell death detection ELISA-kits were purchased from Boehringer 
Mannheim Co. (Indianapolis, IN). 



99 

Experiment 1 

Young purebred (20 months of age; n=36; Angus, An, temperate B. taurus . n=9; 
Romosinuano, Ro, tropically adapted B, taurus, n=9; Brahman, Br, B. indicus , n=9; 
Nelore x Brahman, NB, B. indicus, n=9) bulls were evaluated for testicular and 
spermatogenic traits and extragonadal sperm reserves at the Subtropical Agricultural 
Research Station, Brooksville, Florida (28° 37' N latitude, 82° 22' W longitude) during 
the summer. Animals were sacrificed using standard industry procedures at Central 
Packing Inc. (Center Hill, Florida). Testes were collected and immediately transported on 
ice to the laboratory. Each spermatic cord was trimmed, and the intact testis and tunic 
were weighed. The tunic was then removed, and both the tunic and the testis without 
tunic were weighed separately. Testicular circumference was measured, and testis length, 
depth, and width were obtained to determine testicular volume. Paired testicular volume 
was calculated as the sum of the volume of the right and the left testes. Testes were 
considered paraboloids, and the volume was calculated using the equation v = ui^h, where 
r = (width + depth) / 4 and h = length (Chase, et al., 1997). Epididymides were removed, 
weighed and mid-parenchymal sections were obtained for spermatogenesis quantification 
(5g). 

Quantitative spermatogenesis was assessed by counting elongated spermatids to 
determine daily sperm production (DSP) and DSP/g of testicular parenchyma (DSPG) 
(Chenoweth et al, 1994). Briefly, 5 g of testicular parenchyma were thawed, finely 
minced, and homogenized for approximately 1 min with 25 ml of working solution (0.9% 
NaCl, 0.05% Triton X-100, and 100 ppm Merthiolate diluted 1 :4 with distilled water). 



100 
Then, 175 ml of working solution were added, mixed for 1 min, allowed to settle for at 
least 1 h, and then thoroughly mixed using a stirrer. Number of spermatozoa and 
elongated spermatids were counted using a hemocytometer, and the values were used to 
determine DSPG and DSP using the formula: 

DSPG = AX (B+Y) / (Time divisor)Y 
DSP = DSPG (0.99Z) 
Where X = hemocytometer count, Y = parenchyma sample weight, Z = testis parenchyma 
weight, A = hemocytometer constant, B = dilution factor, and time divisor (TD) (5.32 = 
TD for Bos taurus, An and Ro; and 5.1 1 = TD for Bos indicus , Br and NB) (Amann et al., 
1974; Chenoweth et al., 1994). For extragonadal sperm reserves, the right epididymis 
was separated into caput (head), corpus (body), and cauda (tail). Sperm reserves were 
calculated using the following formula: 

ESR = AX (B+W) 
Where A = hemocytometer constant, X = hemocytometer count, B = dilution factor, and 
W = weight of relevant segment of epididymis (head, body or tail) (Wildeus, 1993). 
Experiment 2 

Tropically adapted young crossbred bulls (n=l 12; averaging 17 months of age; 
Senepol x Angus, SA; Tuli x Angus, TA; Brahman x Angus, BA) were evaluated for 
testicular and spermatogenic traits (as described for experiment 1) and spermatogenic 
apoptosis at the Subtropical Agricultural Research Station, Brooksville, Florida. The 
experiment was repeated over two years using 58 (SA, n=18; TA, n=22; BA, n=18) bulls 
in year 1 and 54 (SA, n=22; TA, n=18; BA, n=14) bulls in year 2. 



101 
Daily sperm production (DSP) and DSPG and in BA were calculated by using 
both the TD for Bos taurus and for Bos indicus (5.32 and 5. 1 1 respectively), whereas only 
the TD for Bos taurus was employed for the other two breeds (SA and TA). 

Aliquots of homogenized testicular parenchyma in DPBS were used for 
quantification of spermatogenic apoptosis using a cell death detection ELISA-kit (Hingst 
and Blottner, 1995) in a subset of 10 bulls/breed/year (n=60). The cell death detection 
ELISA is a quantitative sandwich-enzyme-immunoassay that uses monoclonal mouse 
antibodies against DNA and histones that allows determination of mono- and 
oligonucleosomes in the cytoplasmic fraction of cell lysates. Testicular parenchyma (1 g 
in 2 ml DPBS) was minced, freeze/thawed 3 times, homogenized for 1 min using a 
Polytron (Brinkmann Instruments, Westbury, NY), sonicated for 1 min at 4° C and then 
centrifugation at 25000 g for 30 min at 4° C. The supernatant was collected and stored in 
100 ul aliquots at -20° C. The microtiterplate-modules (MM) were coated with 100 ul of 
coating solution containing the anti-histone antibody, and cultured overnight at 4°C. 
Incubation buffer 200 ul were pipetted into each well, covered with plate cover foils, and 
incubated at room temperature (18° to 25° C) for 30 min to saturate the non-specific 
binding sites of the MM. After incubation, the incubation buffer was removed, and MM 
were washed three times with 300 ul washing solution. Then 100 ul of each sample 
solution containing 10 ug of testis equivalent/ 100 ul were added to the MM, covered, 
and incubated for 90 min at room temperature. For background determination, 100 ul of 
incubation buffer were pipetted into three wells. 



102 
Then, 100 ^1 of anti-DNA-peroxidase were added to the MM and incubated for 90 min at 
room temperature. After incubation, ABTS® (Boehringer) substrate was added to the 
wells, incubated at room temperature for 10 min, and the absorbance read at 405 nm. 
Experiment 3 

A third group of 7 Brahman (Brahman, n=2 and miniature Brahman, n=5, 
averaging 31 and 52 months of age respectively ) bulls as well as 6 Romosinuano (19 
months of age) bulls were analyzed for testicular and spermatogenic traits (DSP and 
DSPG, using a TD=5. 1 1 for Brahman and miniature Brahman and a TD=5.32 for 
Romosinuano) and spermatogenic apoptosis during the summer in Florida as described 
for previous experiments . 
Statistical Analysis 

Data for experiment 1 , 2, and Brahman and miniature Brahman were analyzed by 
least squares ANOVA using SAS (1989 ). In experiment 1, the model included breed, 
and the residual was used as the error term. In experiment 2, the model included year, 
breed, and breed by year. Data was analyzed with and without correcting for body 
weight, and the residual was used as the error term. Preplanned orthogonal contrasts (BA 
vs SA and TA, SA vs TA) were used to compare the effect of breed. In experiment 3, 
Brahman and miniature Brahman model included breed and the residual was used as the 
error term. For Romosinuano, means and standard error of the mean were calculated for 
testicular and spermatogenic traits as well as for spermatogenic apoptosis. 



103 
Results 

Experiment 1 

Angus and Br bulls were older at sacrifice than Ro and NB (P<0.05) Table 5-1. 
Body weight and testicular traits (LSMeans ± SEM for testicular circumference, paired 
testicular volume, paired testicular weight, and paired epididymal weight) in B. taurus 
and B. indicus bulls (An, Br, Ro, NB) are shown in Table 5-1 . Paired testicular volume 
and weight as well as right epididymal head weight were greater in An bulls when 
compared to Br, Ro, and NB (P<0.05). 

Least squares means for spermatogenesis (DSP and DSPG) and extragonadal 
sperm reserves (caput, corpus, and cauda epididymis) in young B. taurus and B. indicus 
bulls (An, Br, Ro, and NB) are shown in Table 5-2. There was no effect of breed on 
DSP, DSPG, ESR (caput), and ESR (cauda). However, ESR (corpus) in Br was higher 
than for An and Ro (3.1 ±0.54 vs 1.2 ±0.55 and 1.7 ± 0.57 xlO 9 spermatozoa 
respectively; P< 0.05). 
Experiment 2 

Least squares means (±SEM) for age, body weight, and testicular traits (paired 
testicular volume, paired testicular and epididymal weight and testicular circumference) 
in crossbred (BA, SA, and TA) bulls pooled for both years are shown in Table 5-3. No 
difference was found in age among breeds, but BA were heavier at sacrifice than SA and 
TA bulls (485.0 ± 8.68 vs 427.5 ± 7.74 and 427.8 ± 7.74 kg; P< 0.05). 



104 



Table 5- 1 . Physical and testicular traits in young Bos taurus and Bos indicus bulls 3 . 

Breed 

Item Angus Brahman Romosinuano Nelore x 

Brahman 

No. of bulls 8 8 8 8 

Age, days bc 662.4 ±8.11 658.6 ±8.11 630.7 ±8.11 634.7 ±8. 11 

Body Weight, kg 489.4 ± 13.23 466.7 ± 13.23 457.6 ± 13.23 473.5 ± 13.23 

Testicular 

circumference, cm 18.0 ±0.72 17.6 ±0.67 18.1 ±0.70 18.1 ±0.68 

Paired testicular 

vol d , cm 3 765.3 ±63.29 600.5 ± 54.47 602.6 ±53.59 570.8 ± 54.85 

Paired testicular 

wt d , g 489.6 ±34.06 388.1 ±31.72 404.3 ±33.72 434.0 ±32.24 

Paired epididymal 

wt, g 50.1 ±3.11 45.5 ±2.90 46.4 ±3.03 45.5 ±2.94 

a Least squares means ± SEM. 

b Contrast, Romosinuano vs Angus and Brahman (P< 0.05). 

c Contrast, Nelore x Brahman vs Angus and Brahman (P< 0.05). 

d Contrast, Angus vs Brahman, Romosinuano, and Nelore x Brahman (P< 0.05). 



105 



Table 5-2. Spermatogenic traits and extragonadal sperm reserves in young Bos taurus 
and Bos indicus bulls 3 . 



Breed 



Item 



Angus 



Brahman 



Romosinuano Nelore x Brahman 



No. of bulls 

Spermatogenesis 

DSP b (x 10 9 ) 

DSPG c (xl0 6 ) 

Epididymal 
Extragonadal 
sperm reserves 

Caput (xlO 9 ) 

Corpus (xlO 9 )* 6 

Cauda (xlO 9 ) 



5.3 ±0.58 
21.9 ±2.02 



3.7 ±0.55 5.4 ±1.73 
19.7 ±1.94 24.1 ±6.07 



4.5 ±0.53 
21.2 ±1.87 



4.9 ±0.89 
1.2 ±0.55 
2.8 ± 0.76 



4.5 ±0.93 3.3 ±0.90 
3.1 ±0.54 1.7 ±0.57 
2.4 ±0.79 3.7 ±0.77 



3.7 ±0.84 
2.6 ±0.55 
3.1 ±0.72 



a Least squares means ± SEM. 

b Daily sperm production, DSP (x 10 9 ). 

c Daily sperm production per gram of testicular parenchyma DSPG (x 10 6 ). 

d Contrast, Brahman vs Angus and Romosinuano (P< 0.05). 



e Contrast, Nelore x Brahman vs Angus (P< 0.05). 



106 



Table 5-3. Physical and testicular traits in young crossbred bulls" 



Breed 


Item Brahman x Angus 


Senepol x Angus 


Tuli x Angus 


No. of bulls 


32 


40 


40 


Age, days 


496 ± 8.7 


511 ± 7.8 


518±7.8 


Body weight 6 , kg 


485 ± 8.7 


428 ± 7.7 


428 ± 7.7 


Testicular 








circumference bce , cm 


17.4 ±0.22 


17.9±0.18 


18.2 ±0.18 


Testicular 








circumference d , cm 


18.1 ±0.25 


17.6 ±0.22 


17.9 ±0.22 


Paired testicular vol bce , 








cm 3 


720 ± 27.0 


734 ±21.8 


777 ±21.8 


Paired testicular 








vol de , cm 3 


799 ± 28.9 


705 ± 25.5 


748 ± 25.5 


Paired testicular wt bce , 








g 


447 ±16.6 


473 ±13.7 


489 ±13.6 


Paired testicular wt d , 








g 


499 ±18.5 


453 ±16.5 


469 ±16.5 


Paired epididymal 
wt b - c - e , g 


51.5 ±1.65 


52.5 ± 1.36 


52.9 ±1.36 


Paired epididymal 

wt d - e , g 


55.1 ±1.67 


51.1 ±1.49 


51.5 ±1.49 


"Least squares means ± SEM. 
b Adjusted for body weight. 
c Adjusted for body weight (P: 
d Unadjusted data. 


sO.01). 







e Orthogonal contrast, Brahman x Angus vs Senepol x Angus and Tuli x Angus (P<0 .05). 



107 
Testicular circumference was lower in BA than in SA and TA (17.4 ± 0.22 vs 17.9 ±0.18 
and 18.2 ± 0.18 cm; P< 0.05). However, breed type did not affect other testicular traits 
evaluated. There was an effect of body weight on testiticular traits (P< 0.01). 

Least squares means (±SEM) for spermatogenic traits (DSP and DSPG) in 
crossbred bulls are shown in Table 5-4. When spermatogenesis was calculated using a 
TD for R taurus (5.32), both DSP and DSPG were lower for BA in comparison with SA 
and TA (5.4 ± 0.38 vs 6.2 ± 0.31 and 6.6 ± 0.31 x 10 6 ; P<0.05 and 1.2 ± 0.12 vs 1.5 ± 
0.10 and 1.6 ± 0.10 x 10 9 ; P<0.05, respectively). In contrast, when spermatogenesis for 
BA alone was calculated using a TD for B. indicus (5.11), only DSP was lower for BA 
when compared with SA and TA (1.3 ± 0.12 vs 1.5 ± 0.10 and 1.6 ± 0.10 x 10 9 ; P<0.05, 
respectively). In both cases, DSP was affected by body weight (P< 0.01). 

Results of estimates of spermatogenic apoptosis, as measured by cell death 
detection ELISA, in crossbred bulls are shown in Figure 5-1. There was no effect of 
breed on spermatogenic apoptosis when BA, SA, and TA were compared (1264 ± 179, 
1039.5 ± 184.23, and 917.6 ± 179.32 mU/mg of testicular parenchyma, respectively). 
Experiment 3 

Least squares means (± SEM) for testicular traits, spermatogenesis, and 
spermatogenic apoptosis in Brahman and Miniature Brahman bulls are shown in Table 5- 
5. Paired testicular volume, paired testicular weight, paired epididymal weight, and DSP 
were smaller in miniature Brahman than in Brahman bulls (P<0.05). 



108 



Table 5-4. Spermatogenic traits and spermatogenic apoptosis in young crossbred bulls 3 







Breed 




Item 


Brahman x Angus 


Senepol x Angus 


Tuli x Angus 


No. of bulls 


32 


40 


40 


Time divisor 


5.32 


5.32 


5.32 


DSP bdeg (xl0 9 ) 


1.2±0.12 


1.5 ±0.10 


1.6±0.10 


DSP b - f (xl0 9 ) 


1.4±0.11 


1.5± 0.10 


1.6±0.10 


DSPG cAg (xl0 6 ) 


5.4 ±0.38 


6.2 ±0.31 


6.6 ±0.31 


DSPG cfg (xl0 6 ) 


5.5 ±0.34 


6.1 ±0.31 


6.6 ±0.31 


Time divisor 


5.11 


5.32 


5.32 


DSP bAeg (xl0 9 ) 


1.3 ± 0.12 


1.5±0.10 


1.6±0.10 


DSP bf (xl0 9 ) 


1.5 ± 0.11 


1.5 ±0.10 


1.6± 0.10 


DSPG cd (xl0 6 ) 


5.6 ±0.39 


6.2 ±0.31 


6.6 ±0.31 


DSPG cf (xl0 6 ) 


5.7 ±0.35 


6.1 ±0.31 


6.6 ±0.31 


No. of bulls 


20 


20 


20 


Spermatogenic 








apoptosis, mU/mg 


1264 ±179.0 


1040 ±184.2 


918 ± 179.3 



a Least squares means ± SEM. 

b Total daily sperm production (DSP x 10 9 ). 

c Daily sperm production per gram of testicular parenchyma (DSPG x 10 6 ). 

d Adjusted for body weight. 

"Adjusted for body weight (P<0.01). 

TJnadjusted data. 

g Orthogonal contrast, Brahman x Angus vs Senepol x Angus and Tuli x Angus (P<0.05). 



109 






E 
E 

(A 

to 
o 

■*-> 

Q. 

O 

Q. 
< 

U 
'c 

03 

a 

o 

TO 

E 
CO 



1400 - 




T 












1200 - 






j 


1000 - 






H 










800 - 








600 - 














400 - 














200 - 














- 




i , 




i 





BA 



SA 
Breed 



TA 



Figure 5-1 . Least squares means (± SEM) for spermatogenic apoptosis in crossbred bulls 
(BA= Brahman x Angus SA= Senepol x Angus, and TA= Tuli x Angus). 



110 



Table 5-5. Testicular traits, spermatogenesis, and spermatogenic apoptosis in Brahman 
and miniature Brahman bulls 3 . 



Breed 



Item 



Brahman 



Miniature Brahman 



No. of bulls 

Age, days 

Testicular circumference, cm 

Paired testicular vol. b , cm 3 

Paired testicular wt b , g 

Paired epididymal wt b , g 

DSP cb (xl0 9 ) 

DSPG d (xl0 6 ) 

Spermatogenic apoptosis, mU/mg 



941.5 ±694.84 

15.1 ±1.15 

557.9 ±15.07 

437.5 ±11.56 

57.6 ±3.35 

2.7 ±0.16 

12.0 ±1.90 

2779.8 ±140.29 



1573.4 ±439.46 

15.3 ±0.94 

402.6 ±12.30 

278.8 ± 9.44 

44.0 ± 2.73 

1.5 ± 0.13 

9.9 ±1.20 

2592.1 ±88.72 



"Least squares Means ± SEM. 

b Brahman was different from Miniature Brahman (P<0.05). 

c Daily sperm production, DSP (x 10 9 ). 

d Daily sperm production per gram, DSPG (x 1 6 ). 



Ill 

In contrast, testicular circumference, DSPG, and spermatogenic apoptosis did not differ 
among breeds. Means ± SEM for testicular traits, spermatogenesis, and spermatogenic 
apoptosis in Romosinuano bulls are shown in table 5-6. Romosinuano bulls in 
experiment 3 were 67 d younger than Ro bulls in experiment 1 . Romosinuano bulls in 
experiment 3 showed similar testicular traits (paired testicular volume, paired testicular 
weight, paired epididymal weight, and testicular circumference) when compared with the 
Ro bulls in experiment 1, although DSP and DSPG were less in Ro in experiment 3 than 
Ro bulls in experiment 1 . 

Discussion 

A comparison of testicular traits, spermatogenesis, and extragonadal sperm 
reserves in 20 mo Angus, Romosinuano, Brahman, and Nelore x Brahman bulls showed 
differences in paired testicular volume and paired testicular weight However, 
quantitative spermatogenic traits (DSP and DSPG) did not differ among temperate 
(Angus) and tropically adapted (Romosinuano, Brahman, and Nelore x Brahman) breeds. 
Extragonadal sperm reserves were not different among breeds with the exception of those 
in the ESR (corpus). When tropically adapted crossbreeds were compared for testicular 
traits, spermatogenesis, and spermatogenic apoptosis, results showed differences in 
testicular circumference and possibly spermatogenesis. However, spermatogenic 
apoptosis was not different among crossbreeds. 



Table 5-6. Testicular traits, spermatogenesis, and spermatogenic apoptosis in 
Romosinuano bulls". 



112 



Breed 



Item 



Romosinuano 



No. of bulls 

Age, days 

Testicular circumference, cm 

Paired testicular vol., cm 3 

Paired testicular wt, g 

Paired epididymal wt, g 

DSP b (xl0 9 ) 

DSPG C (x 10 6 ) 

Spermatogenic apoptosis, mU/mg 






563.0 ±4.22 

17.5 ±0.46 

652.6 ± 46.92 

426.7 ± 24.63 

49.6 ±2.35 
2.7 ±0.32 

12.3 ±1.06 
1730.6 ±193.16 



"Means ± SEM. 

b Daily sperm production, DSP (x 10 9 ). 

c Daily sperm production per gram SPG (x 10 6 ). 



113 
In experiment 1, An and Br bulls were older than Ro and NB by approximately 1 
month. Paired testicular volume and paired testicular weight were larger in An than in 
Br, Ro, and NB. In contrast, BW, testicular circumference, and paired epididymal weight 
were not different among breeds. Angus and Ro (B. taurus) had higher DSP than those 
reported for yearling Angus and Hereford bulls in Colorado which had similar testicular 
weights (Berndtson and Igboeli, 1989). However, contrary to the hypothesized results for 
this study, spermatogenic efficiency (DSPG) did not differ among temperate and 
tropically adapted breeds. As expected, paired testicular weight, ESR (caput), and ESR 
(cauda) in this study, were lower in An and Ro (20 mo of age) than in 7 year old Angus 
and Hereford bulls (Weisgold and Almquist, 1979). Also, testicular traits, DSP, DSPG, 
and ESR for An and Ro ( temperate and tropically adapted B. taurus . respectively) were 
lower than those reported for B. taurus dairy bulls (Amann et al., 1976). However, DSP 
and DSPG were higher, and ESR (corpus) in the present study were similar when 
compared to the results reported by Weisgold and Almquist (1979). 

In the present study, no difference was found between Br and NB in any of the 
parameters studied. Paired testicular and epididymal weights were smaller for Br and NB 
than those reported for Brahman bulls of similar age and body weights in Florida and 
Texas (Chenoweth et al., 1994; Chase et al., 1994). In contrast, paired testicular weight 
was similar to that reported for B. indicus strains in north Australia by Wildeus and 
Entwistle (1982), whereas paired testicular volume was greater for Br and NB in this 
experiment in comparison with previous results from Brahman bulls in Texas (Chase et 
al., 1994). 



114 
Daily sperm production (that is influenced by total testicular parenchyma) was 
less in Br and NB bulls than those reported for Brahman bulls in Florida (Chenoweth et 
al., 1994), but greater than those reported for Brahman bulls in Texas (Rocha et al., 1996) 
and R indicus bulls in tropical Australia (Wildeus and Entwistle, 1982). However, 
DSPG (reflecting spermatogenic efficiency) was greater in Br and NB bulls in this study 
than those reported for R indicus bulls in Florida, Texas, and Tropical Australia 
(Wildeus and Entwistle, 1982; Chenoweth et al., 1994; Rocha et al., 1996). Extragonadal 
sperm reserves (Caput) and ESR (corpus) for Br and NB were larger in our study as 
compared with previous reports for R indicus bulls (Wildeus and Entwistle, 1982; Rocha 
et al., 1996). Extragonadal sperm reserves (cauda) was larger in the present study than 
those reported by Rocha et al. (1996) but smaller than those reported by Wildeus and 
Entwistle (1982). These results are in agreement with a previous report where breed of 
bull did not influence extragonadal sperm reserves (Coulter et al., 1987). However, in 
another study, extragonadal sperm reserves have been reported to be influenced by breed 
and diet (Coulter and Kozub, 1984). 

In experiment 2, although BA bulls were heavier than SA and TA at 508 d of age, 
their testicular circumference was smaller. These results are in agreement with previous 
reports in bulls of similar age. For example, when R indicus bulls were compared for 
development with R taurus (from 269 to 619 d of age), R indicus bulls were heavier 
than R taurus at equivalent ages, but scrotal circumference was smaller (Chase et al., 
1997). This could be due to slower development of scrotal circumference in R indicus 
breeds when compared to R taurus (Chenoweth, et al., 1996). Body weight at 508 d of 



115 
age for these tropically adapted crossbreeds were similar for those reported for 3 year old 
B. indicus x B, taurus crossbred bulls in Australia (Wildeus and Entwistle, 1983). Paired 
testicular weight and paired epididymal weight were greater in all crossbreds evaluated in 
this study than those reported for 3 year old B. indicus x B, taurus crossbred bulls in 
Australia (Wildeus and Entwistle, 1983), whereas DSP and DSPG values were higher in 
B. indicus x B. taurus crossbred bulls reported by Wildeus and Entwistle (1983). 

In the present study, testicular circumference, paired testicular volume, paired 
testicular weight, and paired epididymal weight did not differ among breeds, and were of 
similar magnitude for the same traits reported for Brahman (637 d of age) bulls in another 
study (Chase et al., 1994). 

Daily sperm production/g of testicular parenchyma (DSPG) refers to 
spermatogenic efficiency, whereas DSP is proportional to the total sperm production of 
the total testicular parenchyma. In this study, DSP and DSPG were lower for BA than SA 
and TA when a TD for B. taurus was used. In contrast when a B. indicus TD was used to 
calculate DSP and DSPG for BA only, DSP was lower in BA when compared to SA and 
TA. Daily sperm production/g was similar among breed, indicating that spermatogenic 
efficiency was similar among breeds. However, DSP (and perhaps DSPG) were lower in 
BA bulls (depending on TD used), suggesting that the BA bulls might have had less 
testicular parenchyma than either SA or TA bulls. This was reinforced in this experiment 
where BA had smaller testicular circumferences than the other breed types. Another 
possible explanation for the difference in spermatogenic traits observed for BA bulls 
when compared to SA and TA, could be the R indicus genetic component of BA, since it 



116 
has been reported that B. indicus originated bulls reached puberty later than B. taurus and 
had slower testicular development subsequently than B, taurus breeds (Stewart et al., 
1980; Fields et al., 1982; Chenoweth et al., 1996; Chase et al., 1997). Daily sperm 
production/g and DSP for bulls in experiment 2 were considered to be within values 
reported for B. indicus strain bulls of similar development in Australia (Wildeus and 
Entwistle, 1982), but lower than those reported for yearling Hereford and Angus bulls 
(Coulter et al., 1987; Berndtson and Igboeli, 1989). Also, DSP and DSPG were both 
lower than those reported for Brahman crossbred, Bali cattle ( B. sondaicus ). and hybrid 
bulls ranging from 3 to 7 years of age (N'Dama et al., 1983; Wildeus and Entwistle, 
1983; Cardoso and Godinho, 1985; McCool, 1990; Tegegne et al., 1992). 

In experiment 3, the testicular traits of Brahman bulls were within the ranges 
reported for Brahman bulls in Florida (Chenoweth et al., 1994). Miniature Brahman were 
smaller than Brahman bulls, and the difference in testicular traits (paired testicular 
volume, paired testicular weight, and paired epididymal weight) are probably as a result 
of smaller body size. However, testicular circumference did not differ between Brahman 
and miniature Brahman despite 40% difference in testicular weight. Daily sperm 
production was lower in miniature Brahman than in Brahman, because DSP is influenced 
by total testicular parenchyma, and the testes were smaller in miniature Brahman than in 
Brahman bulls. In contrast, DSPG (spermatogenic efficiency) was not different among 
the two breeds, and since spermatogenic efficiency did not differ, spermatogenic 
apoptosis was also not expected to be different. This indicates that the rate of cellular 



117 
damage within the spermatogenic epithelium caused by summer elevated temperatures in 
Florida is similar among miniature Brahman and Brahman bulls. 

In conclusion, when temperate B. taurus and tropically adapted B, taurus and B, 
indicus bulls of similar age and raised under the same conditions were compared, An 
bulls were heavier and some testicular traits were greater than in tropically adapted bulls 
(Br, Ro, and NB). However, spermatogenesis and extragonadal sperm reserves did not 
differ among breeds, with the exception of ESR (corpus) in Br bulls which was greater 
than in An and Ro bulls. Testicular and spermatogenic traits, including levels of 
spermatogenic apoptosis, in tropically adapted crossbred breeds used in the present study 
(i.e. SA, TA, and BA), were not different when assessed during the Florida summer. In 
contrast, DSP (and perhaps DSPG) were lower in BA than in SA and TA bulls in 
experiment 2. Here, as it is assumed that DSPG was similar among breeds, this would 
indicate that spermatogenic efficiency did not differ among breeds, and the difference in 
DSP was most probably related to differences in the amount of testicular parenchyma as 
reflected in testicular size. Also, more information regarding testicular traits, 
spermatogenesis, and spermatogenic apoptosis was obtained for miniature Brahman as 
compared with Brahman bulls, as well as for the Romosinuano (B. taurus ) bulls during 
the summer in Florida. Brahman and miniature Brahman bulls were older than the rest of 
bulls used in this series of experiments, and their levels of spermatogenic apoptosis were 
higher than levels in younger bulls. Then, it is possible to speculate that age could 
influence the levels of spermatogenic apoptosis, resulting in higher levels of 
spermatogenic apoptosis in older than in younger bulls. 



CHAPTER 6 
SUMMARY AND CONCLUSIONS 



The effect of elevated temperatures and gossypol toxicity on spermatogenic 
apoptosis, testicular traits, and the use of antioxidants to counter the effects of gossypol in 
bulls were studied, as well as breed differences (B. indicus and B. taurus, tropically 
adapted vs B. taurus temperate) in response to elevated temperatures. 

In Chapter 3, two experiments were designed to induce and assess spermatogenic 
apoptosis. Experiment 1 , tested the hypothesis that spermatogenic stress, caused by 
elevation of body temperature, by E. coli endotoxin or elevation of testicular temperature, 
by scrotal insulation, increases spermatogenic apoptosis. In this experiment three groups 
were included; 1) control (n=3), 2) R, coli endotoxin (100 ng/kg) (n=4), and 3) scrotal 
insulation for 48 h (n=4). Bulls were sacrificed 10 days after initiation of treatment. 
Results of experiment 1, showed an effect of time of assessment on respiration rate and 
rectal temperature(P<0.05 respectively). However, there was no apparent effect of 
treatment on testicular and semen traits, spermatogenic traits determined by DSP and 
DSPG, and level of spermatogenic apoptosis. 

In experiment 2, the hypothesis that SI increased spermatogenic apoptosis was 
tested, with additional objectives being to test breed differences in spermatogenic 
apoptosis and changes in levels of spermatogenic apoptosis with respect to time after SI. 



118 



119 
This experiment included two groups; 1) control (Angus, temperate B. taurus, n = 4; 
Senepol, tropically adapted B. taurus , n - 2; Romosinuano, tropically adapted B. taurus, n 
= 2) and 2) scrotal insulation (SI) for 48 h (Angus, n = 4; Senepol, n = 4; Romosinuano, n 
= 2). Bulls were sacrificed at either 2 d or 4 d after initiation of treatments. In 
experiment 2, the body weight of Angus bulls was significantly lower than Senepol and 
Romosinuano (P< 0.001), whereas rectal and scrotal temperatures were higher in Angus 
when compared to Senepol and Romosinuano bulls (P<0.05). However, rectal 
temperature was not affected by scrotal insulation. Scrotal temperature was not affected 
by breed, and the temperature of the scrotum was higher in scrotal insulated after 24 h 
than in control bulls (PO.01). Semen characteristics and paired epididymal weights were 
not affected by 48 h of scrotal insulation or breed. Scrotal circumference, paired 
testicular volume, testicular circumference, and paired testicular weight were smaller in 
Angus bulls than Senepol and Romosinuano (PO.001 respectively). Scrotal insulation 
had no effect on DSPG and spermatogenic apoptosis, although DSP was significantly 
lower in Angus than in Senepol and Romosinuano bulls (PO.01). This suggested that 
spermatogenic efficiency was not different between breeds and the difference in DSP was 
due to differences in testicular size. 

Experiments designed to induce and assess spermatogenic apoptosis in Chapter 3 
failed to detect increases in spermatogenic apoptosis as assessed by histology, tunel stain, 
and cell death detection ELISA in bulls after E. coli endotoxin infusion or 48 h of scrotal 
insulation at 2 d, 4 d, or 10 d after initiation of treatment. This suggested that either these 
treatments caused no spermatogenic apoptosis changes or that the detection windows 






120 
used were inappropriate, since no other signs of cell death were observed when evaluated 
by histology. Also, these results suggest that the levels of spermatogenic apoptosis that 
were observed, correspond to basal levels of spermatogenic apoptosis, resulting probably 
from spontaneous degeneration of germ cells, that undergo apoptosis during normal 
spermatogenesis, (Bianco-Rodriguez and Martinez-Garcia, 1996). It is clear that more 
work should be done in bulls to elucidate the role of spermatogenic apoptosis in 
spermatogenic dysfunction. 

It has been suggested that the deleterious effect of elevated temperatures on 
spermatogenesis occurs through an imbalance or excess of reactive oxygen species 
(Loven et al., 1988). In Chapter 4, an experiment was designed to test the effects of long 
term feeding gossypol on semen traits, testicular traits, spermatogenesis, and 
spermatogenic apoptosis with provision to test this theory via use of a dietary antioxidant 
(vitamin E) in an attempt to counteract spermatoxic effects of gossypol in the bull 
(Velasquez-Pereira, 1995). Gossypol is a toxic phenolic pigment found in the cotton 
plant, and cotton products containing gossypol such as cottonseed meal are commonly 
used as feedstuffs for cattle. In this experiment young Holstein bulls (n=24; 6 mo of age) 
were distributed into three experimental groups (each n=8); 1) CONT, supplemented with 
soybean meal (SBM), corn, and 30 IU of vitamin E/kg, 2) GOSS, supplemented with 
cotton seed meal (CSM), corn, and 30 IU vitamin E/kg, and 3) G+VITE, supplemented 
with, corn, and 4,000 IU vitamin E/bull/d. Supplements GOSS and G+VITE were 
formulated to supply 14 mg of free gossypol/kg body weight (BW)/d, and bulls were 
supplemented from 6 to 15 mo of age. 



121 
Results showed that the percentage of normal spermatozoa was lower and 
percentage of primary sperm abnormalities was higher in the GOSS supplemented bulls 
when compared to CONT or G+VITE (P<0.05). In contrast, no treatment effect was 
observed on secondary sperm abnormalities, paired testicular and epididymal weight, and 
scrotal circumference. Spermatogenesis, described in terms of DSP and DSPG, was 
lower (P<0.05) in gossypol fed bulls compared with either control bulls (CONT) or those 
fed both gossypol and 4000 IU of vitamin E (G+VITE). Surprisingly, gossypol had no 
effect on spermatogenic apoptosis, despite significantly affecting sperm production, 
although there was an inverse relationship between sperm production (both DSP and 
DSPG) and the level of spermatogenic apoptosis (r=-0.55; P<0.05 and r=-0.46; PO.05, 
for DSP and DSPG respectively). Less than 1 apoptotic cell per seminiferous tubule was 
observed (tunel stain or histology) in the spermatogenic epithelium in all three groups 
(CONT, GOSS, and G+VITE). 

Free gossypol had an adverse effect on percentage of normal sperm, percentage of 
primary abnormalities, DSP, and DSPG, although testicular measures and spermatogenic 
apoptosis were not affected. Vitamin E fed at 4,000 IU vitamin E/animal/d was able to 
block the spermatoxic effects of gossypol on spermatogenesis, suggesting that vitamin E 
supplementation in the diets of bulls fed with cottonseed meal could allow this valuable 
feedstuff to be fed safely to intact bulls. 

In Chapter 5, the relative susceptibility of B. taurus and B. indicus and crossbred 
bulls to environmentally elevated temperature effects on testicular and spermatogenic 
traits and spermatogenic apoptosis were evaluated during the summer in central Florida. 



122 
Two experiments were designed to test the hypothesis that tropically adapted B, taurus 
and B. indicus breeds will have better semen and testicular traits and lower 
spermatogenic apoptosis than a temperate breed (Angus) during the summer. In 
experiment 1, young B. taurus and B, indicus bulls (20 months of age; n=36; Angus, 
temperate B. taurus . An, n=9; Romosinuano, tropically adapted B. taurus , Ro, n=9; 
Brahman, tropically adapted B. indicus , Br, n=9; Nelore x Brahman, tropically adapted B, 
indicus, NB, n=9) were evaluated for testicular and spermatogenic traits and extragonadal 
sperm reserves at the Subtropical Agricultural Research Station, (28° 37' N latitude, 82° 
22' W longitude), Brooksville, Florida. Results for this experiment showed that paired 
testicular volume and paired testicular weight were greater in An bulls when compared to 
Br, Ro, and NB (P<0.05). There was no effect of breed on DSP, DSPG, ESR (caput), and 
ESR (cauda). However, ESR (corpus) in Br was higher than An and Ro (P< 0.05). 
In experiment 2, the objective was to compare tropically adapted B. taurus 
crossbred (Senepol x Angus and Tuli x Angus) with tropically adapted B. taurus x B. 
indicus crossbred (Brahman x Angus). Young crossbred (17 months of age; n=l 12; 
Senepol x Angus, SA; Tuli x Angus, TA; Brahman x Angus, BA) bulls were evaluated in 
June for testicular and spermatogenic traits and spermatogenic apoptosis at the 
Subtropical Agricultural Research Station, Brooksville, Florida. The experiment was 
performed twice using 58 (SA, n=18; TA, n=22; BA, n=18) bulls in year 1 and 54 (SA, 
n=22; TA, n=18; BA, n=14) in year 2. Results showed that bull ages were similar. 
Brahman x Angus bulls were heavier at sacrifice than SA and TA bulls (PO.05). 
Testicular circumference was lower in BA than in SA and TA (P< 0.05). However, breed 



123 
did not affect other testicular traits evaluated. When spermatogenesis was calculated 
using a TD for R taurus (5.32), DSP and DSPG for BA were lower than for SA and TA 
(P<0.05). In contrast, when spermatogenesis for BA was calculated using a TD for B, 
indicus (5.11) while the other breeds used the B. taurus TD, DSP was significantly lower 
when compared to SA and TA (P<0.05). In this experiment, DSP and perhaps DSPG 
were lower in BA than SA and TA bulls. Daily sperm production/g was similar among 
breeds, and the difference on DSP was as a result of smaller testicular size in BA. 
Spermatogenic apoptosis was not affected by breed type. 

A third group of 7 Brahman (Brahman, n=2 and miniature Brahman, n=7) bulls as 
well as 6 Romosinuano bulls were evaluated for testicular traits, spermatogenic traits 
(DSP and DSPG, using a TD=5.1 1 for Brahman and miniature Brahman and a TD=5.32 
for Romosinuano), and spermatogenic apoptosis. This information was of interest 
because there is little information on reproduction traits for these breeds. Paired testicular 
volume, paired testicular weight, paired epididymal weight, and DSP were smaller in 
miniature Brahman than in Brahman bulls (P<0.05). In contrast, testicular circumference, 
DSPG, and spermatogenic apoptosis did not differ among breeds. 

Romosinuano bulls in experiment 3 were 67 d younger than Romosinuano bulls in 
experiment 1 . Romosinuano bulls in experiment 3 showed similar testicular traits (paired 
testicular volume, paired testicular weight, paired epididymal weight, and testicular 
circumference) when compared with Romosinuano bulls in experiment 1 , although DSP 
and DSPG were less in Romosinuano in experiment 3 than for Romosinuano bulls in 
experiment 1. 



124 
In Chapter 5, when testicular traits and extragonadal sperm reserves (ESR) were 
assessed during the Florida summer in R taurus and R indicus bulls, An bulls were 
heavier and some testicular traits were greater when compared to Br, Ro, and NB. 
However, spermatogenesis and epididymal sperm reserves were not different among 
breeds, with the exception of ESR (corpus) in Br bulls. When the crossbred breeds used 
in the present study (BA, SA, and TA) were compared during the florida summer, 
testicular traits and spermatogenic apoptosis did not differ among breed types. In 
contrast, DSP and perhaps DSPG, were found to be lower in BA bulls when both were 
calculated using a TD for B. taurus . In contrast, when a TD for B, indicus was used for 
BA bulls only, DSP was lower (P^ 0.05) in BA bulls compare to SA and TA bulls. Also, 
more information regarding testicular traits, spermatogenesis, and spermatogenic 
apoptosis was obtained for miniature Brahman as compared with Brahman bulls, as well 
as for the tropically adapted Romosinuano (R taurus ) bulls during the summer in Florida. 
Brahman and miniature Brahman bulls were considerably older than the rest of bulls used 
in this series of experiments, and the levels of spermatogenic apoptosis in these bulls 
were considerably higher than the levels observed in younger animals in this study, 
suggesting that age could influence the levels of spermatogenic apoptosis. 

In conclusion, experiments in this study failed to observe increases in 
spermatogenic apoptosis in the bull as a result of E. coli endotoxin infusion, scrotal 
insulation (48 h), gossypol toxicity, or summer temperatures in central Florida. Also, 
breed type did not appear to influence the levels of spermatogenic apoptosis when 
tropically adapted and temperate breeds were compared. The reasons for these findings 



125 
could include 1) inappropriate detection windows used in experiments in Chapter 3, 2) 
none of the treatments used in this series of studies caused an increased in spermatogenic 
apoptosis, 3) germ cell death resulting from elevated temperatures or gossypol toxicity 
might take a different form of cell death than spermatogenic apoptosis, 4) an age 
influence on levels of spermatogenic apoptosis, resulting in higher levels in older than in 
younger bulls, making detection of small changes in spermatogenic apoptosis in young 
bulls difficult. 

Gossypol increased sperm abnormalities and decreased spermatogenic traits (DSP 
and DSPG), but gossypol showed no effect on spermatogenic apoptosis. The antioxidant 
vitamin E fed at 4,000 IU vitamin E/bull/d counteracted the spermatoxic effect of 
gossypol in all parameters evaluated in this study, suggesting that vitamin E 
supplementation when feeding cottonseed meal could allow the safe use of this valuable 
feedstuff with intact bulls. 

Overall this study failed to demonstrate differences in semen and testicular traits 
among crossbred or B. taurus and B. indicus (tropically adapted vs temperate) bulls 
compared in Chapter 5. Spermatogenic efficiency per gram of testicular parenchyma 
(DSPG) was lower in BA B. indicus x B. taurus crossbred bulls (Chapter 5, experiment 
1) when a TD for B. taurus was used to calculate DSPG for BA only. In contrast, if a TD 
for B. indicus was used DSPG did not differ. 

This series of studies suggests that future work should be oriented to identify an 
optimal time to measure spermatogenic apoptosis after a heat insult such as scrotal 
insulation for 48 h. These experiments should include a positive control group where 



126 
scrotal insulated (48 h) bulls should be kept and ejaculated every 3 days to ensure that the 
scrotal insulation treatment causes the spermatogenic damage that is expected. Also, 
semen should be collected in all bulls to measure semen markers for spermatogenic 
apoptosis such as Hsp70 level. In this experiment, spermatogenic apoptosis should be 
assessed (by histology, by tunel stain, by DNA laddering, and by cell detection ELISA, as 
well as by new technologies) daily until d 16 after initiation of scrotal insulation (0 d). 
Spermatogenic apoptosis results should be correlated with sperm morphology obtained in 
the positive control group, as well as with semen markers such as levels of Hsp70 in bull 
semen. 



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BIOGRAPHICAL SKETCH 

Victor Hugo Monterroso Perez was born October 1, 1965, in Guatemala City, 
Guatemala. He is the third son of Nehemias Monterroso Salvatierra and Zoila Lidia 
Perez Mazariegos. He was raised in Guatemala City and in the town of San Pedro 
Yepocapa. He received his bachelor's diploma in sciences from the "Adolfo V. Hall" 
Military Institute of the Guatemalan Army in 1982, and later he was commissioned as an 
Infantry Lieutenant. Victor Monterroso was subsequently admitted to the College of 
Veterinary Medicine at the University of San Carlos of Guatemala. He finished his 
course work in 1989 and in the same year married his wife, Norma I. Mora, and moved to 
Miami, Florida. In Spring 1991, Victor enrolled at University of Florida as an 
undergraduate student of the Dairy Science Department where he had the opportunity to 
work under the supervision of Dr. Peter J. Hansen. The journal article "Regulation of 
bovine and ovine lymphocyte proliferation by progesterone: modulation by steroid 
receptor antagonist and physiological status" published in Acta Endocrinologica (1993) 
was a product of his undergraduate research. In December 1992, he received his 
Bachelor of Science degree from the University of Florida, and in Spring 1993 he 
enrolled as a graduate student in the Department of Dairy Science. In March 1 1 of the 
same year his son Victor Alejandro was born, making him a blessed father. In August 
1995, he received his Master of Science degree from the University of Florida. 

148 



149 
Upon completion of the Master of Science degree in 1995, he completed all the 
requirements to obtained the title of Medico Veterinario at the degree of Licenciado 
(veterinarian) from the University of San Carlos of Guatemala in his native country of 
Guatemala. At the same time he began his doctoral degree in Veterinary Medical 
Sciences at the College of Veterinary Medicine of the University of Florida where he is 
now a candidate for the degree of Doctor of Philosophy. In 1997, he was awarded the 
Florida Veterinary Medical Association Auxiliary Graduate Studies Scholarship. In that 
year, he became the President of the Veterinary Graduate Student Association and the 
college representative of the University of Florida Graduate Student Council. In 1998, he 
was awarded the Auxiliary Graduate Studies Scholarship. After completion of his 
studies, he is planning to pursue either a training in laboratory animal medicine, a training 
in food animal medicine, a reproductive physiology postdoctoral position, or return to his 
native Guatemala. 



I certify that I have read this study and that in my opinion it conforms to 
acceptable standards of scholarly presentation and is fully adequate, in scope and quality, 
as a dissertation for the degree of Doctor of Philosophy. 




' 





Peter J. (Chenoweth, Chair 
Associate Professor of 
Veterinary Medicine 

I certify that I have read this study and that in my opinion it conforms to 
acceptable standards of scholarly presentation and is fully adequate, in scope and quality, 
as a dissertation for the degree of Doctor of Philosophy. 




Louis F. Archbalc 
Professor of Veterinary 
Medicine 

I certify that I have read this study and that in my opinion it conforms to 
acceptable standards of scholarly presentation and is fully adequate, in scope and quality, 
as a dissertation for the degree of Doctor of Philosophy. 

£^o 2). {jUl±? 

Claus D. Buergelt ^ 
Professor of Veterinary 
Medicine 

I certify that I have read this study and that in my opinion it conforms to 
acceptable standards of scholarly presentation and is fully adequate, in scope and quality, 
as a dissertation for the degree of Doctor of Philosophy. 




Chadwick C. Chase, Jr. 
Assistant Professor of 
Animal Science 

I certify that I have read this study and that in my opinion it conforms to 
acceptable standards of scholarly presentation and is fully adequate, in scope and quality, 
as a dissertation for the degree of Doctor of Philosophy. 





^enneth C. Drury 
Clinical Associate Profess 
Obstetrics/Gynecology 



This dissertation was submitted to the Graduate Faculty of the College of 
Veterinary Medicine and to the Graduate School and was accepted as partial fulfillment 
of the requirements for the degree of Doctor of Philosophy. 



December, 1998 

Dean, College of Veterina 
Medicine 




Dean, Graduate School